University of Nebraska - Lincoln
DigitalCommons@University of Nebraska - Lincoln
Dissertations and Theses in Biological Sciences
Biological Sciences, School of
Spring 4-8-2016
No Nonsense: The Protection of Wild-Type
mRNAs From Nonsense-Mediated mRNA Decay
in Saccharomyces cerevisiae
Krista Patefield
University of Nebraska-Lincoln, [email protected]
Follow this and additional works at: http://digitalcommons.unl.edu/bioscidiss
Part of the Biology Commons, and the Other Microbiology Commons
Patefield, Krista, "No Nonsense: The Protection of Wild-Type mRNAs From Nonsense-Mediated mRNA Decay in Saccharomyces
cerevisiae" (2016). Dissertations and Theses in Biological Sciences. Paper 79.
http://digitalcommons.unl.edu/bioscidiss/79
This Article is brought to you for free and open access by the Biological Sciences, School of at DigitalCommons@University of Nebraska - Lincoln. It
has been accepted for inclusion in Dissertations and Theses in Biological Sciences by an authorized administrator of DigitalCommons@University of
Nebraska - Lincoln.
No Nonsense: The Protection of Wild-Type mRNAs From Nonsense-Mediated mRNA
Decay in Saccharomyces cerevisiae
by
Krista D. Patefield
A DISSERTATION
Presented to the Faculty of
The Graduate College at the University of Nebraska
In Partial Fulfillment of Requirements
For the Degree of Doctor of Philosophy
Major: Biological Sciences
(Genetics, Cellular and Molecular Biology)
Under the Supervision of Professor Audrey L. Atkin
Lincoln, Nebraska
April, 2016
No Nonsense: The Protection of Wild-Type mRNAs From Nonsense-Mediated mRNA
Decay in Saccharomyces cerevisiae
Krista D. Patefield, Ph.D.
University of Nebraska, 2016
Advisor: Audrey L. Atkin
Gene regulation in eukaryotes is tightly controlled at multiple levels to ensure
proper expression and cellular homeostasis. Misregulation of gene expression is a
common source of genetic disease. One mechanism by which cells are able to control
gene expression is through the synthesis and degradation of the mRNA molecules
encoding the genes. The transcription and degradation of mRNA molecules controls the
pool mRNAs that are available to the translational machinery. One of the well-studied
mRNA decay pathways is the Nonsense-Mediated mRNA Decay pathway (NMD).
Originally, NMD was discovered as a posttranscriptional mRNA surveillance mechanism
responsible for the deadenylation-independent decapping and rapid 5’3’ degradation of
mRNAs that harbor premature termination codons (PTCs). Approximately one-third of
all inherited genetic disease and cancers are related to NMD. It is now known that NMD
plays a much larger role in the stability and expression of wild-type mRNAs as well.
Wild-type mRNAs with NMD-targeting signals, which include 1) a translated uORF, 2) a
long 3’ UTR, 3) leaky scanning leading to out-of-frame initiation of translation, 3)
programmed ribosome frameshift sites, and 5) regulated alternative splicing variants, are
rapidly destabilized by NMD. It has also been observed that some wild-type mRNAs
contain NMD targeting signals but are not degraded by NMD due to protecting
mechanism. Here we show that the SSY5 mRNA in Saccharomyces cerevisiae is a wild-
type mRNA with multiple NMD targeting signals but is not degraded by NMD. None of
the current models for NMD protection explain the SSY5 mRNA stability so the
mechanism of protection is likely to be novel. Additionally, we show the SSY5 mRNA is
primarily degraded 5’3’. We also explore two additional mRNAs, YAP1 and GCN4, in
S. cerevisiae that also contain at least one NMD-targeting signal but are not degraded by
NMD. Elucidating the mechanism of protection from NMD of these three mRNAs will
provide valuable insight to the underlying molecular mechanisms of NMD, which despite
thorough investigation remain unclear. Understanding the molecular intricacies of the
NMD pathway will allow for the efficient development of NMD-related disease therapies
with minimal risks and side-effects.
Graphical Abstract
v
ACKNOWLEDGEMENTS
This trip through graduate school has been unlike anything I have ever
experienced. There is no way I would have made it through to the end without the
guidance, support and encouragement of a multitude of people. First of all, I would like
to thank my advisor Dr. Audrey Atkin. You took a risk by allowing me to join your lab
with very little prior research experience. You were patient with my learning curve and
provided me with all of the tools I needed to be successful. I was lucky to have joined a
lab that encouraged not only quality research but also emphasized mentoring, teaching,
and work-life balance. It has been a privilege to spend the past 6 ¾ years learning from
you.
Next, I would like to thank the members of my wonderful committee: Dr. Ken
Nickerson, Dr. Mark Wilson, Dr. Steve Harris, and Dr. Jeff Mower. You have truly been
a great committee to work with and have never made me feel uncomfortable during
exams or committee meetings. Although my project is distant from most of your areas of
work, you have always listened diligently and offered great advice. At times scheduling
meetings was tricky but you all were very kind in working with me so that I could meet
requirements and deadlines. I really could not have asked for a better, kinder group of
faculty to work with.
I would also like to extend a sincere thank you to my undergraduate advisor and
friend from Hastings College, Dr. John Bohmfalk. Dr. B., you are the reason I made it
here in the first place! Thank you for believing in me and encouraging me to give this a
shot. The scientific foundation you provided me with—both in the classroom and in the
vi
lab—was a significant reason for my smooth transition into graduate school. Thank you
for challenging me and teaching me how to study, learn, and apply my knowledge.
There are also some faculty and support staff here at UNL that played a
significant role in my success. Dr. Valery Forbes, although you are no longer here at
UNL it was truly a privilege to work with you. From my very first encounter with you
requesting funds and support for the Biology Graduate Student Symposium, as well as
requesting a key note address from you for the symposium, you have been so kind and
accommodating. Thank you for being such a great leadership example. Tammy Kortum,
you have always been there to answer questions and sort things out. You were wonderful
to work with during the planning of the Biology Graduate Student Symposium. Mindy
Peck, thank you so much for answering all of my questions, for bailing me out of
paperwork mishaps, and for letting me vent about frustrating days—you have been
wonderful. Alan Muthersbaugh and Les Acree, thank you so much for doing everything
you could to always get me the supplies I needed as soon as you could and find the best
deals on products. I know you have bent over backwards and gone out of your way for
me so that my work could keep running smoothly and your efforts were very much
appreciated.
To all of my friends and lab members (past and current), thank you for the
support, encouragement, advice, and brainstorming sessions. Michelle Palmer, Jessica
Hargarten, Ruvini Pathirana, Nur Ras Aini Ahmad Hussin, Hannah O’Neill, Maya
Khasin, Esther Choi, Melanie Langford, Sahar Hasim, Swetha Tati, Xinrong Ma,
Raghuveer Singh, Derrick White, Yukari Maezato, Annabel Olson, Anna Lampe, and
vii
Jeff Bunker, I could write a page for each one of you thanking you for the multiple things
you have done to help me out. To my friends outside of the lab, Jen Endres and Homero
Flores, Christa and David Serpa, Bonnie and Will Ratcliff, Deb Calhoun, Matt and Katie
Meyer, and Sally Schedlock, I appreciate your support, encouragement and
understanding more than you could possibly know.
Finally, I want to thank my family. I truly would have never made it through this
journey without your continued loving support. Mom and dad, words cannot describe my
gratitude for supporting me in every way possible. You have given me everything I
could have ever dreamed of and so much more. I am truly blessed to be your daughter.
To my in-laws, Judy, Dan, Megan, Mike, Travis, and all of the extended family, thank
you for encouraging and supporting in me over the years. I couldn’t have married into a
better family. And to my husband, Jason—the one who has been by my side every day of
this journey. Every time I doubted myself you encouraged me and maintained
unwavering faith in me. You believed in me when I didn’t believe in myself. Thank you
for helping to cook, clean, do laundry, yard work, and handling the day-to-day life tasks
that I didn’t have time for. You made so many sacrifices for us, for me, so that I could
pursue this. I’ll never be able to truly thank you enough for your love and support.
And last, I want to close by extending a thank you to our family members that are
no longer with us. Jason and I have lost many very important people to us during my
time in graduate school and I know they are cheering me on from above. Josephine
Fager, Virgil Fager, Wallace Shaw, Milo Patefield, Paul Guern, Colleen Snyder, Sheryl
Marchant and Margaret Hammond—we love and miss you! Thank you for everything. ♥
viii
TABLE OF CONTENTS
ACKNOWLEDGEMENTS ..............................................................................................v
TABLE OF CONTENTS .............................................................................................. viii
LIST OF FIGURES ......................................................................................................... xi
LIST OF TABLES ......................................................................................................... xiii
CHAPTER 1: Introduction ..............................................................................................1
References ..........................................................................................................................31
CHAPTER 2: Saccharomyces cerevisiae SSY5 mRNA is a wild-type mRNA with
multiple NMD-targeting signals but is not degraded by NMD ..................................46
Abstract ..............................................................................................................................47
Introduction ........................................................................................................................48
Methods..............................................................................................................................51
Results ................................................................................................................................57
Discussion ..........................................................................................................................81
Figure Legends...................................................................................................................86
Figures................................................................................................................................96
Tables ...............................................................................................................................109
References ........................................................................................................................110
ix
CHAPTER 3: SSY5 mRNA decay is predominantly 5’ to 3’ ....................................118
Abstract ............................................................................................................................119
Introduction ......................................................................................................................120
Methods............................................................................................................................124
Results ..............................................................................................................................127
Discussion ........................................................................................................................138
Figure Legends.................................................................................................................141
Figures..............................................................................................................................145
Tables ...............................................................................................................................151
References ........................................................................................................................152
CHAPTER 4: The protection of the Saccharomyces cerevisiae wild-type GCN4 and
YAP1 mRNAs from degradation by NMD ..................................................................158
Abstract ............................................................................................................................159
Introduction ......................................................................................................................160
Methods............................................................................................................................163
Results ..............................................................................................................................166
Discussion ........................................................................................................................181
Figure Legends.................................................................................................................187
Figures..............................................................................................................................192
Tables ...............................................................................................................................199
References ........................................................................................................................200
x
CHAPTER 5: N5-Phosphonoacetyl-L-ornithine (PALO): influence on regulation of
amino acid biosynthetic genes in Saccharomyces cerevisiae ......................................206
Abstract ............................................................................................................................207
Introduction ......................................................................................................................208
Methods............................................................................................................................208
Results ..............................................................................................................................210
Discussion ........................................................................................................................213
Figure Legend ..................................................................................................................214
Figure ...............................................................................................................................216
References ........................................................................................................................217
CHAPTER 6: Summary and Future Directions ........................................................218
References ........................................................................................................................231
xi
LIST OF FIGURES
Figure 2-1 ...........................................................................................................................96
Figure 2-2 ...........................................................................................................................97
Figure 2-3 ..................................................................................................................... 98-99
Figure 2-4 .........................................................................................................................100
Figure 2-5 .........................................................................................................................101
Figure 2-6 ................................................................................................................. 102-103
Figure 2-7 .........................................................................................................................104
Figure 2-8 .........................................................................................................................105
Figure 2-9 .........................................................................................................................106
Figure 2-10 .......................................................................................................................107
Figure 2-11 .......................................................................................................................108
Figure 3-1 .........................................................................................................................145
Figure 3-2 .........................................................................................................................146
Figure 3-3 .........................................................................................................................147
Figure 3-4 .........................................................................................................................148
Figure 3-5 .........................................................................................................................149
Figure 3-6 .........................................................................................................................150
Figure 4-1 .........................................................................................................................192
Figure 4-2 .........................................................................................................................193
Figure 4-3 .........................................................................................................................194
Figure 4-4 .........................................................................................................................195
Figure 4-5 .........................................................................................................................196
xii
Figure 4-6 .........................................................................................................................197
Figure 4-7 .........................................................................................................................198
Figure 5-1 .........................................................................................................................216
xiii
LIST OF TABLES
Table 2-1 ..........................................................................................................................109
Table 3-1 ..........................................................................................................................151
Table 4-1 ..........................................................................................................................199
1
CHAPTER 1:
Introduction
2
1.0 Overview
Eukaryotic gene expression is an incredibly intricate process and is controlled
with high fidelity at many different levels. When something goes awry at any level in the
tight control of gene expression the result can be detrimental not only to the cell but to
the organism as a whole manifesting in cell death and disease. The most obvious level of
control is at the level of protein synthesis from the mRNA transcript (translation).
However, a level of control even before that of translation occurs with the biogenesis of
the mRNA transcripts (transcription) and also turnover of the mRNA transcripts
(degradation). The processes of mRNA transcription and degradation are also tightly
controlled. Eukaryotes have evolved several quality control (QC) mechanisms to help
maintain the fidelity of gene expression by quickly riding the cell of aberrant mRNAs
(Lykke-Andersen and Bennett, 2014). It is now known that these pathways can also be
exploited to regulate gene expression of wild-type mRNAs. One of the most extensively
studied QC mechanisms is the nonsense-mediated mRNA decay (NMD) pathway.
However, despite extensive study and characterization over the past several years the
exact molecular mechanisms of NMD are not entirely known.
The NMD pathway was originally characterized for its role in the rapid
degradation of mRNAs that contain a premature termination codon (PTC). If these PTCcontaining mRNAs were not rapidly removed from the translatable pool they would lead
to the build-up of C-terminally truncated proteins, which could result in deleterious
consequences (Akimitsu, 2008; Baker and Parker, 2004; Behm-Ansmant et al., 2007;
Chang et al., 2007; Frischmeyer and Dietz, 1999; Hentze and Kulozik, 1999; Hilleren and
3
Parker, 1999; Muhlemann et al., 2008; Schweingruber et al., 2013; Shyu et al., 2008).
The NMD pathway plays a vital role in the regulation of gene expression and exists in all
eukaryotes that have been examined including yeast, Drosophila, Caenorhabditis
elegans, Arabidopsis, and humans (Bedwell et al., 1997; Grimson et al., 2004; Hall and
Thein, 1994; He et al., 2003; Hentze and Kulozik, 1999; Kalyna et al., 2012; Kurihara et
al., 2009; Maquat and Carmichael, 2001; Maquat and Serin, 2001; Mendell et al., 2004;
Pulak and Anderson, 1993; Rayson et al., 2012; Rehwinkel et al., 2005; Sun and Maquat,
2000). Importantly, it is now recognized that the NMD pathway is also responsible for
the degradation of a significant portion of wild-type mRNAs (non-PTC-containing) as
well. Studies in Saccharomyces cerevisiae, Drosophila, C. elegans, Arabidopsis, and
humans have revealed that a significant portion of the transcriptome is affected when the
NMD pathway is inactivated (Guan et al., 2006; He et al., 2003; Johansson et al., 2007;
Kalyna et al., 2012; Lelivelt and Culbertson, 1999; Mendell et al., 2004; Mitrovich and
Anderson, 2005; Peccarelli and Kebaara, 2014; Rayson et al., 2012; Rehwinkel et al.,
2005).
The NMD pathway requires to coordinated action of the three core trans-acting
factors, up-frameshift proteins Upf1, Upf2 and Upf3, which have orthologs in all
eukaryotes examined (Applequist et al., 1997; Cali et al., 1999; Cui et al., 1995; Denning
et al., 2001; Gatfield et al., 2003; He and Jacobson, 1995; Hodgkin et al., 1989; Isken and
Maquat, 2008; Jeong et al., 2011; Leeds et al., 1991; Leeds et al., 1992; Lykke-Andersen
et al., 2000; Pulak and Anderson, 1993). It has been shown that mutation or deletion in
any of the genes coding for these three factors results in the stabilization of NMD-
4
substrates (Cui et al., 1995; He and Jacobson, 1995; Leeds et al., 1991; He et al., 1997;
Lee and Culbertson, 1995; Lelivelt and Culbertson, 1999; Maderazo et al., 2000). Upf1
is the most highly conserved of the three NMD factors and is enriched in binding to
NMD substrates (Johansson et al., 2007). This ~109 kDa protein exhibits both RNAdependent ATPase activity and 5’3’ ATP-dependent RNA helicase activity in addition
to RNA binding (Altamura et al., 1992; Bhattacharya et al., 2000; Czaplinski et al., 1995;
Weng et al., 1996a, b). Upf1 interacts with the eukaryotic translation release factors
eRF1 and eRF3, and upon recognition of an NMD substrate interacts with Upf2 (directly)
and Upf3 (indirectly) to form a surveillance complex (Czaplinski et al., 1999; Czaplinski
et al., 1998; Ivanov et al., 2008; Kadlec et al., 2006). There is evidence to support the
idea that the formation of the surveillance complex occurs through the staged assembly of
two subcomplexes—1) Upf1/eRF1/eRF3 and 2) Upf2/Upf3—which come together to
form a complete NMD mRNP (Atkin et al., 1997). Additionally, Upf1 has also been
shown to interact with mRNA decay factors Dcp1 and Dcp2 as well as Ski7, an exosomeassociated protein (Lykke-Andersen, 2002; Takahashi et al., 2003). Upf1 localizes
predominantly to the cytoplasm, but can shuttle between the nucleus and the cytoplasm
(Lykke-Andersen et al., 2000; Mendell et al., 2002). Upf2 interacts with both Upf1 and
Upf3 likely serving as a bridge between the two proteins (He et al., 1997; LykkeAndersen et al., 2000). The N-terminal domain of Upf2 contains multiple nuclear
localization signals but the protein localizes predominantly to the perinuclear region of
the cytoplasm (Lykke-Andersen et al., 2000; Mendell et al., 2002; Muhlemann et al.,
2008). Upf3 is the smallest and least conserved of the three core NMD factors and
5
contains multiple nuclear localization and nuclear export signals (Lee and Culbertson,
1995; Lykke-Andersen et al., 2000; Serin et al., 2001; Shirley et al., 1998).
The exact mechanism by which the cellular machinery identifies an NMDsubstrate, whether the mRNA contains a PTC or is a wild-type mRNA with an NMDtargeting signal (discussed below), among all mRNAs remains unclear despite thorough
investigation. However, it is known that targeting of an mRNA for degradation by NMD
begins with the recruitment and assembly of the NMD mRNP, which consists of the three
core factors discussed above (Nicholson et al., 2010; Rebbapragada and Lykke-Andersen,
2009). Upf1is the first to interact with the terminating ribosome and release factors eRF1
and eRF3. Upf1 then binds to the Upf2/Upf3 subcomplex, mRNA decay factors are
recruited, and mRNA decay is initiated through deadenylation-independent decapping by
the Dcp1/Dcp2 decapping complex and 5’3’ mRNA decay by the exonuclease Xrn1
(Coller and Parker, 2004; Decker and Parker, 1993; Hsu and Stevens, 1993; Maderazo et
al., 2000; Muhlrad et al., 1994, 1995; Nicholson et al., 2010; Rebbapragada and LykkeAndersen, 2009). It has been shown that presence of poly(A)-binding protein (Pab1),
which is bound to the 3’ poly(A) tail of the mRNA, is able to inhibit assembly of the
NMD mRNP when in close proximity to the terminating ribosome (Amrani et al., 2004;
Behm-Ansmant et al., 2007; Beilharz and Preiss, 2007; Gallie, 1991; Silva et al., 2008;
Wilusz et al., 2001). However, the presence of Pab1 is not absolutely required for the
recognition of NMD substrates by the cellular machinery (Meaux et al., 2008; Roque et
al., 2015). Additionally, some mRNAs with long 3’ UTRs—which distances the
interaction between the terminating ribosome and Pab1—are not affected by NMD-
6
mediated mRNA degradation (Kebaara and Atkin, 2009; Nicholson et al., 2010;
Obenoskey et al., 2014; Rebbapragada and Lykke-Andersen, 2009, Chapter 2). These
observations suggests that other mechanisms exist that are able to antagonize activation
of the NMD pathway. Because of the critical role of NMD in the regulation of gene
expression and in the development of many human diseases it is critical in moving
forward to identify the exact mechanism(s) by which the cellular machinery distinguishes
an NMD substrate from the rest of the mRNA pool.
2.0 NMD in Human Health
Due to the vital role of NMD in the fidelity of gene expression it is not surprising
that as many as 33% of all genetic disorders and inherited cancers are directly linked to
NMD (Culbertson, 1999; Frischmeyer and Dietz, 1999). The majority of these result as a
consequence of the cell not being able to produce sufficient levels of full-length protein.
A few of these prominent disorders include β-thalassemia, Duchene’s and Becker’s
Muscular Dystrophy, Marfan Syndrome, and Cystic Fibrosis (Bhuvanagiri et al., 2010;
Frischmeyer and Dietz, 1999; Khajavi et al., 2006).
β-thalassemia, a disorder of hemoglobin production, is an example that portrays
the protective effects of NMD. This disease has multiple genetic and phenotypic variants
all related to NMD and PTCs in the HBB mRNA which codes for the β-globin protein. A
homozygous PTC mutation able to trigger NMD in both β-globin alleles results in a
deficiency of tetrameric hemoglobin, which requires both α-globin and β-globin, and
leads to severe anemia. However, individuals that are heterozygous for the NMD-
7
competent PTC in the β-globin gene are usually able to produce sufficient β-globin and
remain physically healthy. Both of these cases, in which the PTC elicits NMD, result in a
recessive mode of inheritance. In contrast, a dominant negative form of β-thalassemia
arises when an individual is heterozygous for an NMD-incompetent PTC (Hall and
Thein, 1994; Thein et al., 1990a; Thein et al., 1990b). In this case the PTC does not elicit
NMD due to a positional effect of its location in the last exon (exon 3) of the β-globin
gene (Nagy and Maquat, 1998; Thermann et al., 1998). One model of NMD in
mammalian cells posits that in order to trigger NMD in mammalian cells a PTC must be
at least 50-55 nt upstream of the final exon junction that is marked by an exon-junction
complex (EJC; Neu-Yilik et al., 2011; Thermann et al., 1998). Because this mutant βglobin gene does not trigger NMD it results in the production of toxic levels C-terminally
truncated β-globin which accumulates in precipitation bodies (Neu-Yilik et al., 2011;
Peixeiro et al., 2011). These individuals are severely anemic and sometimes require
transfusions to survive among other developmental complications (Cao and Galanello,
2010).
Marfan Syndrome is a connective tissue disorder that results from mutations in
fibrillin 1 mRNA, and is another example of the protective effects of NMD. Nonsense
mutations in the fibrillin 1 mRNA that trigger NMD leading to significantly reduced
accumulation of functional protein are associated with a milder disease phenotype.
However, when mutant mRNA escapes degradation by NMD and truncated protein is
allowed to build up the disease phenotype becomes much more severe (Dietz, 1993;
Dietz et al., 1993).
8
The other side of NMD in human disease is the aggravation of disease severity
due to haploinsufficiency because of the degradation of mRNAs that could potentially
produce proteins with partial function. Duchenne’s muscular dystrophy (DMD) is a
severe form of muscular dystrophy that is caused by lack of functional dystrophin protein
in skeletal and cardiac muscles. As many as 98% of the mutations in the dystrophin open
reading frame result in the introduction of a PTC and subsequent elimination of the
mRNA by NMD. However, a less severe form of muscular dystrophy, Becker’s
muscular dystrophy (BMD), results when mRNAs with PTCs are able to escape NMD
and produce partially functionally C-terminally truncated dystrophin protein (Kerr et al.,
2001).
NMD also plays a role in a portion of Cystic Fibrosis patients as well. Cystic
fibrosis results from mutations in the mRNA that codes for Cystic Fibrosis
Transmembrane Conductance Regulator (CFTR) protein. Several different mutations in
the mRNA can result in the clinical manifestation of Cystic Fibrosis. Different mutations
in the mRNA are categorized into classes and affect the CFTR protein at different stages
of development. Class I mutations are those that result in a PTC and cause the
degradation of the mRNA through the NMD pathway. Thus, the CFTR protein is never
able to reach the cell membrane where it is needed. These mutations represent
approximately 10% of the mutations that cause Cystic Fibrosis (Bhuvanagiri et al., 2010;
Rogan et al., 2011).
9
2.1 Nonsense Codon Readthrough Therapies
Given the role of NMD in the clinical outcome of a significant portion of
detrimental diseases it is of no surprise the NMD pathway is a targeted area of clinical
research. As discussed above, NMD-associated disease occurs when 1) mRNAs with
PTCs are rapidly removed from the translatable pool of mRNAs so necessary levels of
functional protein are never translated, 2) mRNAs with PTCs are able to avoid
degradation by NMD but still harbor the PTC resulting in the production of only partially
functional protein, or 3) mRNAs containing PTCs escape NMD which leads to the buildup of toxic levels of truncated protein.
Different therapies are being investigated based on two classes of nonsense
mutations: 1) those that introduce a PTC as the result of point mutations so the rest of the
mRNA can still be translated in the native reading frame, and 2) those that introduce a
PTC as the result of a frameshift mutation leaving the remainder of the mRNA to be
translated in an alternate reading frame. The end goal, which is subject to overcoming
many hurdles, is the synthesis of full-length functional protein from the mutant mRNA
without perturbing the translation of any other mRNAs.
Some of the therapies currently being investigated are the use of aminoglycosides,
small molecule drugs that promote ribosome readthrough at nonsense codons (e.g.
Ataluren), suppressor tRNAs, targeted gene repair, antisense oligonucleotides (AOs), and
the upregulation of proteins that could compensate for loss of the functional protein
(Kuzmiak and Maquat, 2006; Peltz et al., 2013). The latter two of these approaches are
being investigated specifically for the nonsense codons that introduce a PTC as a result of
10
a frameshift mutation, as these provide the more challenging case for therapy
development.
These therapies are promising—some have even made it through late-stage
clinical trials—and provide hope for individuals living with NMD-associated debilitating
diseases. However, there are still many questions related to the basic underlying
mechanism of the NMD pathway that remain unanswered, which makes therapy
development both risky and challenging. Elucidating the basic molecular mechanism of
NMD and how the cell is able to distinguish NMD substrates from non-NMD substrates
will be provide crucial information for the development of nonsense therapies as well as
help to understand the potential risks and side effects of new and current therapies.
3.0 Wild-Type mRNA Degradation by NMD
One way by which we can continue to untangle the underlying mechanism of the
NMD pathway is by studying the category of wild-type mRNAs that are regulated by
NMD. Several wild-type mRNA NMD-targeting signals have already been described.
These include: 1) a long 3’ UTR (Amrani et al., 2004; Kebaara and Atkin, 2009; Muhlrad
and Parker, 1999), 2) translation of an upstream open reading frame (uORF; Amrani et
al., 2006; Barbosa et al., 2013; Gaba et al., 2005; Nyiko et al., 2009), 3) a start codon in a
suboptimal context which can lead to leaky scanning and out of frame initiation of
translation (Welch and Jacobson, 1999), 4) the presence of programmed ribosome
frameshift (PRF) sites (Plant et al., 2004) and 5) the presence of pre-mRNA introns and
11
regulated alternative splicing resulting in PTCs (He et al., 1993; Lewis et al., 2003;
McGlincy and Smith, 2008; Ni et al., 2007).
3.1 NMD-targeting through the presence of a translated uORF
Many mRNAs contain one or more short open reading frames in the 5’ leader
region of the mRNA known as a uORFs (Nagalakshmi et al., 2008). These uORFs can
have many functions but they are best known for their role in regulating mRNA
translation and mRNA turnover (Arribere and Gilbert, 2013). A genome-wide study
conducted in S. cerevisiae projected that out of 220 predicted direct NMD substrates 135
(~61%) contain putative uORFs (Guan et al., 2006). The presence of a uORF, when
translated, leads to the introduction of a stop codon that is far upstream of the native ORF
stop codon and, thus, may be recognized as a PTC targeting the mRNA for degradation
by NMD. It has been shown that active translation of the uORF is required to elicit NMD
while those mRNAs that contain untranslated uORFs are immune to NMD (Hurt et al.,
2013).
At least three mRNAs in S. cerevisiae have been verified as wild-type mRNAs
that are targeted to NMD by a uORF: the CPA1, FZF1, and ALR1 mRNAs (Gaba et al.,
2005; Guan et al., 2006; Johansson and Jacobson, 2010). The FZF1 mRNA, which
encodes a transcription factor involved in sulfite metabolism, provides an interesting case
in which the uORF begins in the 5’ leader and ends in the coding ORF (Breitwieser et al.,
1993; Guan et al., 2006). The uORF has two potential start codons at -64 and -58 and the
stop codon appears in the ORF at position +29. Interestingly, when each of the two
12
uORF start codons was mutated (ATGAGG) individually the FZF1 mRNA remained a
target for NMD. However, when both uORF start codons were mutated simultaneously
the FZF1 mRNA sensitivity to NMD was abolished. This data indicates that the uORF,
as long as both uORF start codons are able to be utilized, is an NMD-targeting feature for
the FZF1 mRNA (Guan et al., 2006).
The CPA1 mRNA encodes the small subunit of carbamoyl phosphate synthetase,
which plays a role in the synthesis of citrulline—an arginine precursor (Pierard et al.,
1979). The CPA1 mRNA was originally identified as a wild-type mRNA that is sensitive
to NMD (He et al., 2003; Ruiz-Echevarria and Peltz, 2000) and was later shown to be
targeted to NMD by presence of the uORF (Gaba et al., 2005). The study by Gaba et al.,
(2005) showed that mutation of the uORF start codon (AUGUUG) in a luciferase
reporter resulted in a 2-fold increase in steady-state accumulation compared to the
reporter with the wild-type uORF AUG codon in NMD+ cells. Additionally, the reporter
with the wild-type uORF AUG codon showed a ~7-fold increase in accumulation in an
nmd- strain compared to the NMD+ strain (Gaba et al., 2005). These results provide
evidence that the CPA1 mRNA is targeted to NMD by presence of the uORF in the 5’
leader.
Another important example of the regulation of an mRNA by NMD in response to
the presence of a uORF is the case of the ALR1 mRNA. The ALR1 mRNA codes for a
magnesium (Mg2+) transporter at the plasma membrane and is responsible for magnesium
homeostasis in yeast (Graschopf et al., 2001; MacDiarmid and Gardner, 1998). A study
by Johansson and Jacobson in 2010 showed that yeast are able to control the fidelity of
13
translation, in part, by controlling magnesium uptake. The control of magnesium uptake
is regulated by the product of the ALR1 mRNA, which is in turn regulated by NMD. The
regulation of the ALR1 mRNA by NMD was shown to depend on the presence of a uORF
in the 5’ leader of the mRNA. The ALR1 mRNA actually contains three uORFs (and one
of which has both an A and a B form), but mutational analysis revealed that NMDsensitivity could be incurred by the 3’-most uORF (uORF3) alone (Johansson and
Jacobson, 2010).
The idea of regulation of mRNA susceptibility to NMD by the presence of a
uORF is not exclusive to S. cerevisiae. Studies in Saccharomyces pombe, C. elegans,
mammals and plants all show evidence of uORF-containing mRNAs that are upregulated
when NMD is inactivated (Calvo et al., 2009; Kochetov et al., 2002; Mendell et al., 2004;
Nyiko et al., 2009; Ramani et al., 2009).
3.2 NMD-targeting through the presence of PRF sites
Programmed ribosome frameshift (PRF) sites are cis-acting mRNA sequences
that cause a ribosome to shift reading frames by one nucleotide either upstream (5’
direction) or downstream (3’ direction), -1 or +1 PRF site, respectively. A -1 PRF site
appears to be the most common phenomenon and is the best characterized of the two.
The -1 PRF site consists of a “slippery site,” which is a heptameric mRNA sequence of
the form X XXY YYZ (in the original reading frame where X is any three identical
nucleotides, Y is either a triplet A or triplet U sequence, and Z is an A, U or C), followed
downstream by an mRNA secondary structure of two or more stem-loops known as a
14
pseudoknot (Baranov et al., 2002; Jacks and Varmus, 1985; Namy et al., 2006). The
slippery site and the pseudoknot are separated by a region of 5-9 nucleotides known as
the “spacer” region (Dinman et al., 1991). The pseudoknot structure causes the ribosome
to pause and shift its reading frame on the slippery site (Namy et al., 2006). The binding
strength of the nucleotides that compose the pseudoknot is directly correlated with the
efficiency at which the structure is able to induce ribosomal frameshifting (Hansen et al.,
2007). However, if the strength of the pseudoknot is too strong then it can actually
decrease the efficiency of translation downstream (Tholstrup et al., 2012). The +1 PRF
site, which is less common, consists of a slippery site and at least one additional
stimulatory element such as a pseudoknot (Ivanov et al., 2000).
In recent years it has become increasingly evident that both +1 and -1 PRF sites
are being used to control gene expression in a variety of organisms (Dinman, 2012;
Namy et al., 2004). It has been well-studied that viruses use -1 PRF sites to synthesize
polyproteins from polycistronic mRNAs as a method of genome consolidation (Baril et
al., 2003; Brierley and Dos Ramos, 2006; Dulude et al., 2002; Jacks and Varmus, 1985;
Leger et al., 2007). Additionally, PRF sites are responsible for the generation of an
Escherichia coli DNA polymerase III subunit (Blinkowa and Walker, 1990) and release
factor 2 (RF2; Baranov et al., 2002), two yeast telomerase-associated proteins (Aigner et
al., 2000; Morris and Lundblad, 1997), actin-binding protein Abp140 in yeast (Asakura et
al., 1998), and all metazoan ornithine decarboxylase antizymes (Ivanov et al., 2000).
A computational study identified the -1 PRF motif at 260 locations in the S. cerevisiae
genome, which is ~5.2-fold greater frequency than random chance (Hammell et al.,
15
1999). Further analysis indicated that many of the S. cerevisiae -1 PRF sites would result
in the ribosome encountering a PTC after resuming translation following the frameshift
event (Plant et al., 2004). In a study by Plant et al., in 2004 the yeast PGK1 mRNA,
which is an extensively characterized and very stable wild-type mRNA, was destabilized
by the addition of a -1 PRF site. This destabilization was NMD-dependent as the
construct regained stability in an nmd- strain. Additional confirmation was done to
confirm that the destabilization in NMD+ strains was the result of a -1 PRF event (Plant
et al., 2004). This same study also showed that there in an inverse correlation between
mRNA stability (half-life) and the efficiency of ribosome frameshifting. Further, steadystate mRNA accumulation is drastically reduced when -1 PRF efficiency is high (Plant et
al., 2004).
Thus, the frequency of PRF events among all organisms is greater than originally
perceived and PRF events could potentially be used as a gene regulatory mechanism
(Hammell et al., 1999; Plant et al., 2004). PRF events that cause the ribosome to
encounter a PTC can subject the mRNA to rapid degradation by NMD which limits the
translational pool of the mRNA.
3.3 NMD-targeting through leaky scanning
The sequence context that surrounds an AUG start codon is largely responsible
for the ability of a ribosome to properly initiate translation at a particular AUG. The -6 to
+6 sequence surrounding the AUG is important in determining translation initiation
efficiency, with the strongest influence at -3 position (Cigan et al., 1988; Yun et al.,
16
1996). The optimal start codon initiation consensus sequence appears to be
(A/U)A(A/C)AA(A/C)AUGUC(U/C) (Hamilton et al., 1987). Should the surrounding
sequence of the AUG codon be suboptimal the ribosome can bypass the AUG in search
of an AUG in a better context. This concept is referred to as “leaky scanning.” If the
ribosome initiates at a different downstream start codon the initiation may occur in an
alternate reading frame, which is referred to as out-of-frame initiation of translation.
Leaky scanning resulting in out-of-frame initiation of translation may lead to the
introduction of a PTC in the alternate reading frame being translated, which would
destabilize the mRNA as a consequence of NMD.
In 1987 Sharp and Li developed a mathematical model, called the codon
adaptation index (CAI), which measured the bias of synonymous amino acid codon
(different codons that code for the same amino acid) usage which had been shown to
have a nonrandom distribution (Sharp and Li, 1987). Later, the CAI equation was
adapted to model start codon context resulting in the AUGCAI equation to show that there
is a relationship between codon usage bias and translation initiation context (Miyasaka,
1999). Finally, the AUGCAI equation was revised into the AUGCAI(r) equation, which was
used to test the leaky scanning model for NMD (Guan et al., 2006). This equation takes
into consideration the weighted contribution of each nucleotide at each position (from -6
to +6) and how frequently each nucleotide is used at each position. The result is a value
from 0 to 1 that measures the likelihood of translation initiation, and consequently the
likelihood of leaky scanning, to occur at a particular AUG with 0 being poor and 1 being
efficient (Guan et al., 2006). Further, limited experimental analysis led to the general
17
conclusion that AUG codons with an AUGCAI(r) score of ≤0.32 might be good candidates
for leaky scanning while those with a score ≥0.44 would be poor candidates for leaky
scanning (Guan et al., 2006). No examples were shown that have scores in the 0.32-0.44
range, so whether AUG codons with values in that range are subject to leaky scanning
remains to be determined.
The SPT10 mRNA is wild-type mRNA in yeast that exhibits sensitivity to NMD.
The SPT10 mRNA does contain a uORF in the 5’ leader region, however, mutational
analysis of the uORF start codon (AUGCCC) revealed that translation of this uORF
has little to do with the susceptibility of the mRNA to NMD (Welch and Jacobson, 1999).
To determine the cis-element responsible for the SPT10 mRNA susceptibility to NMD
chimeric reporters were constructed which contained portions of the SPT10 mRNA and
portions of the well-characterized and stable PGK1 mRNA. These experiments indicated
that the mRNA destabilizing element was located within the 5’ UTR and first 94
nucleotides of the ORF of the SPT10 mRNA. Further analysis revealed that the ORF
AUG of the SPT10 mRNA is in a suboptimal context with an AUGCAI(r) value of 0.32
(Guan et al., 2006). Given this it was confirmed that the ribosome is able to bypass the
first AUG and initiate at an AUG in a better context downstream in the +1 reading frame.
When the ribosome initiates at the second AUG a PTC is encountered just 15 nucleotides
downstream which targets the mRNA for degradation by NMD (Welch and Jacobson,
1999).
18
3.4 NMD-targeting through the presence of a long 3’ UTR
It has also been shown that wild-type mRNAs with long 3’ UTRs are substrates
for degradation by NMD in S. cerevisiae, C. elegans, Drosophila, plants and mammals.
Studies report mRNAs that contain mutations resulting in a long 3’ UTR and synthetic
mRNAs that terminate at a normal stop codon but contain a long 3’ UTR are both
substrates for NMD (Amrani et al., 2004; Muhlrad and Parker, 1999).
This has been most extensively studied in S. cerevisiae in which the normal 3’
UTR length is 50-200 nucleotides with a median length of 121 nucleotides (Graber et al.,
1999). A screen of wild-type mRNAs in S. cerevisiae revealed that 56 mRNAs were
predicted to have a 3’ UTR of longer than 350 nucleotides (an arbitrary cut-off). Of these
56 mRNAs 11 were experimentally tested. Of these 11 mRNAs 10 accumulated to
significantly higher levels in a upf1Δ strain compared to wild-type. Additionally, halflives were determined for 5 of the 11 mRNAs and all 5 tested had a significantly longer
half-life in the upf1Δ strain compared to wild-type (Kebaara and Atkin, 2009). The one
anomaly was the SSY5 mRNA.
The presence of a long 3’ UTR has also been identified as a cis-acting NMDtargeting element in Nicotiana benthamiana and Arabidopsis (Kalyna et al., 2012;
Kertesz et al., 2006). In plants, increasing the length of the 3’ UTR correlates with an
increase in NMD efficiency (Kertesz et al., 2006). The same correlation has been
observed in murine embryonic stem cells as well (Hurt et al., 2013). Additionally, human
mRNAs with long 3’ UTRs are also subject to NMD and some of these mRNAs are those
that code for crucial NMD factors, which predicts a feedback loop (Longman et al., 2013;
19
Yepiskoposyan et al., 2011). Further studies show that this regulatory feedback loop is
conserved across different organisms (Longman et al., 2013).
The collective data from many of these studies supports a model developed by
Amrani et al., in 2004 known as the faux 3’ UTR model. This model is based on the idea
that the cellular machinery recognizes a PTC because the terminating ribosome and
associated factors are positioned too far upstream from the poly(A) tail (Amrani et al.,
2004; Muhlrad and Parker, 1999). The distance between the termination complex and the
poly(A) tail fails to initiate the normal interaction of the ribosome-bound eRF3 with
poly(A) tail-bound protein Pab1 (Behm-Ansmant et al., 2007). Because this interaction
fails to occur the termination complex is subject to recruitment of the NMD factors and
subsequent degradation (Amrani et al., 2004; Brogna and Wen, 2009). Further, artificial
tethering of Pab1 in close proximity to a prematurely terminating ribosome is able to
abrogate NMD-sensitivity (Amrani et al., 2004). The faux 3’ UTR model is able to
explain the degradation of mRNAs by NMD that contain PTCs and also those mRNAs
that are substrates for NMD because of a long 3’ UTR.
This regulation of wild-type mRNAs by NMD due to the presence of a long 3’
UTR may also be physiologically relevant as revealed by the PGA1 mRNA. The PGA1
mRNA codes for an essential component of the GPI-mannosyl-transferase II, which is
involved in the synthesis of GPI anchors that are added to proteins for membrane
attachment (Sato et al., 2007). The PGA1 mRNA also has a 3’ UTR of ~750nt and is a
substrate for NMD (Kebaara and Atkin, 2009). Interestingly, NMD has also been shown
to regulate the expression of a set of mRNAs involved the assembly of GPI-anchored cell
20
wall proteins (Guan et al., 2006). Consistent with this, cells of the upf1Δ strains are much
more sensitive to Calcofluor White (a fluorescent stain that binds to cellulose and chitin
in cells walls and can compromise cell wall integrity at high concentrations; Kebaara and
Atkin, 2009).
3.5 NMD-targeting through pre-mRNA introns and alternative splicing
Wild-type pre-mRNAs that escape to the cytoplasm still containing introns as
well as regulated alternative splicing are related events that also provide wild-type
mRNAs that become NMD substrates. In yeast, the CYH2, RP51B, and MER2 premRNAs are inefficiently spliced and are substrates for the NMD pathway. In wild-type
cells a very small accumulation of all three pre-mRNAs is observed while in upf1Δ cells
the pre-mRNAs accumulate to significantly higher levels and have longer half-lives (He
et al., 1993). It was also shown that the three mRNAs associate with ribosomes, which is
consistent with 1) these inefficiently spliced mRNAs are escaping to the cytoplasm, and
2) the notion that NMD is a translation-dependent mechanism (He et al., 1993; Peltz et
al., 1993). The inefficient splicing of these pre-mRNAs and their subsequent association
with ribosomes results in the translating ribosome, which begins at the normal ORF
translation initiation site, terminating at a PTC, which triggers NMD.
Similarly, many alternative splicing events result in mRNAs that introduce a PTC
in the normal reading frame. Because these alternative splicing events are regulated it is
likely that alternative-splicing induced NMD serves an intentional role in gene regulation.
Studies of regulated alternative splicing events showed that this is indeed the case in
21
mammalian cells where NMD is exploited to achieve post-transcriptional regulation by
destabilizing certain mRNA isoforms (Cuccurese et al., 2005; Lejeune et al., 2001;
Sureau et al., 2001; Weischenfeldt et al., 2012; Wollerton et al., 2004). Notably, the
polypyrimidine tract binding protein, PTBP1, and the SC35 protein are regulators of
alternative splicing and the proteins themselves are encoded by mRNAs that are subject
to alternative-splicing induced NMD in HeLa cells (Sureau et al., 2001; Wollerton et al.,
2004). Alternative-splicing induced NMD is also responsible for the regulation
ribosomal genes in mammalian cells and in C. elegans (Cuccurese et al., 2005; Mitrovich
and Anderson, 2000).
Although a very small percentage of S. cerevisiae mRNAs contain introns, a
recent study by Kawashima and colleagues in 2014 found that of those mRNAs that do
contain introns many produce nonproductive alternatively spliced mRNAs that resulted in
degradation by NMD (Kawashima et al., 2014). Studies also confirm alternatively
spliced mRNA isoforms that are subject to NMD in Drosophila, Arabidopsis and
zebrafish (Hansen et al., 2009; Jaillon et al., 2008; Kertesz et al., 2006).
4.0 Mechanisms of Protection of Wild-Type mRNAs from NMD
Although it has been widely shown that wild-type mRNAs containing certain
NMD-targeting features (and there are likely additional features that remain to be
discovered) are substrates for NMD, there are exceptions. Some mRNAs have one or
more NMD-targeting element discussed above but show no significant difference in
steady-state accumulation or half-life in wild-type versus upf1Δ cells (Kebaara and Atkin,
22
2009; Obenoskey et al., 2014; Ruiz-Echevarria et al., 1998; Vilela et al., 1998) (Chapter
2). A few mechanisms that are able to protect mRNAs with NMD-targeting signals from
degradation by NMD have been discovered. Given that these mechanisms are not able to
explain the stability of all mRNAs that are protected from NMD (e.g. SSY5 mRNA)
future studies are likely to expand this list. It is important that we identify all of the wildtype mRNA NMD-targeting mechanisms as well as the features that are able to protect an
mRNA from degradation by NMD so that we can gain a deeper more comprehensive
understanding of the molecular mechanisms of gene regulation by NMD. This
information can then be exploited for safer and more efficient therapy development for
individuals that are affected by NMD-related diseases.
4.1 Inhibition of translation
Several observations have led to the idea that NMD is a translation-dependent
process. First, NMD is perturbed by translation elongation inhibitors such as
cycloheximide (Herrick et al., 1990; Zhang et al., 1997; Zuk and Jacobson, 1998).
Additionally, NMD is able to resume once the drug is removed (Zhang et al., 1997).
Second, mutations that block translation elongation, such as the addition of strong stemloops (pseudoknots) or mutations in the tRNA nucleotidyltransferase, also inhibit NMD
(Peltz et al., 1992; Tholstrup et al., 2012). Third, PTC-containing mRNAs do cofractionate with polyribosomes in a pattern that reflects the position of the PTC (i.e. early
PTCs result in mRNAs in lighter fractions whereas later PTCs result in mRNAs in the
heavier fractions; He et al., 1993). Fourth, PTC suppression or stop codon readthrough
23
promoted by suppressor tRNAs results in the stabilization of nonsense containing
mRNAs (Gozalbo and Hohmann, 1990; Keeling et al., 2004). Fifth, eukaryotic release
factors eRF1 and eRF3 are essential for NMD and only interact with a terminating
ribosome when a stop codon is recognized in the A site during translation (Bertram et al.,
2001; Czaplinski et al., 1998; Kisselev et al., 2003; Rospert et al., 2005). Together these
observations lead us to the conclusion that if an mRNA is not actively translated then the
mRNA is not likely to be degraded by the NMD machinery.
4.2 Stop Codon Readthrough
Translation termination begins when any of the three stop codons (UAA, UGA
and UAG) move into the ribosomal A site. When the stop codon appears in the A site a
release factor binds since there is no tRNA that corresponds to any of the three stop
codons. Binding of the release factor mediates the cleavage of the polypeptide chain
from the tRNA in the ribosomal P site. In eukaryotes the two release factors are eRF1
(encoded by the SUP45 mRNA) and eRF3 (encoded by the SUP35 mRNA). eRF1, a
class I release factor, recognizes all three stop codons and eRF3, a class II release factor,
forms a complex with eRF1 to mediate termination through GTP-dependent hydrolysis
(Bertram et al., 2001; Kisselev et al., 2003; Rospert et al., 2005).
Just as the start codon of an mRNA has an optimal and suboptimal surrounding
sequence context that varies translation initiation efficiency, the efficiency at which the
stop codon is recognized is also influenced by the surrounding sequence context. In S.
cerevisiae the nucleotide with the most prominent influence is the one immediately
24
following (3’) the trinucleotide stop codon. The nucleotide at this position can influence
the ability of the ribosome efficiently terminate at the stop codon by at least 100-fold
(Bonetti et al., 1995). This has led to the idea that translation termination is actually
directed by a tetranucleotide sequence rather than a trinucleotide sequence. Additionally,
the six nucleotides upstream and downstream of the stop codon can also influence
termination efficiency, but to a smaller degree (Namy et al., 2001). A ribosome can fail
to terminate translation at a termination codon by incorporating a near-cognate
aminoacyl-tRNA, which complements the sequence of the stop codon at two out of three
nucleotide positions, into the ribosomal A site instead of the release factor (Fearon et al.,
1994). This occurrence is known as stop codon suppression, also frequently referred to
as translational readthrough or leaky termination. The result is a continually growing Cterminally extended polypeptide chain that can have significantly altered function, even if
only a few amino acids are added (DePristo et al., 2005; Namy et al., 2002). The ORF
stop codon of many yeast mRNAs is followed immediately by more in-frame stopcodons which can serve as a secondary protection if readthrough of the first stop codon
were to occur (Williams et al., 2004).
It is interesting, but not surprising, that the ability of the ribosome to readthrough
a PTC is able to suppress degradation of the mRNA by NMD. Reading through a PTC
may cause the incorporation of one wrong amino acid, but the rest of the polypeptide
remains unchanged since the reading frame was not altered (Keeling et al., 2004). Once
the ribosome bypasses the PTC by incorporating a near-cognate aminoacyl-tRNA instead
of a release factor translation is able to proceed until the normal ORF stop codon is
25
recognized. This is the basis for the nonsense suppression therapies that target
readthrough of PTC so the cells can produce full-length functional protein (Finkel, 2010;
Gunn et al., 2014; Peltz et al., 2013). A potential challenge to this approach is the ability
to cause readthrough of the PTC without perturbing translation termination at the natural
stop codon, especially if the PTC and natural Stop codon are the same triplet codon.
The idea of stop codon readthrough can also be applied to those mRNAs that are targeted
for degradation by a long 3’ UTR. If the faux 3’ UTR model is correct, which there is
much evidence to support this, then translation termination at a normal stop codon
followed by a long 3’ UTR would place the terminating ribosome in an improper special
context for the termination complex to interact with Pab1 (Amrani et al., 2004; BehmAnsmant et al., 2007; Brogna and Wen, 2009; Muhlrad and Parker, 1999). Thus, if
readthrough of the stop codon were to occur—even though the stop codon is not a PTC—
and terminate at another in frame stop codon downstream (i.e. in closer proximity to the
poly(A) tail), it would result in two important outcomes: 1) a C-terminally extended
polypeptide which could have altered function, either harmful or beneficial, and 2) the
mRNA, which originally had a long 3’ UTR, would no longer be a substrate for NMD.
4.3 Translation Reinitiation
Another mechanism by which mRNAs can avoid degradation by NMD is by
reinitiation of translation at a downstream AUG after termination at a PTC. Evidence of
reinitiation as an NMD-antagonist comes from a studies of the human β-globin mRNA.
In humans, NMD appears to be positional dependent. It has been shown that typically
26
mRNAs with PTCs at the 3’ end of the mRNA fail to trigger NMD which can result in
the build-up of C-terminally truncated proteins. This follows an exon-junction complex
(EJC) model in which presence of an EJC triggers degradation of an mRNA if
termination occurs more than 50-55nt upstream of the final EJC (Chamieh et al., 2008;
Le Hir et al., 2001; Lykke-Andersen et al., 2001; Shibuya et al., 2004; Singh et al., 2007).
EJC-independent NMD has also been observed, but most of these examples are based on
the improper spatial context of the terminating ribosome and the poly(A) tail. This
follows the faux 3’ UTR model in that PTCs located near the 3’ end of the mRNA, thus
in closer spatial context to the poly(A) tail, may fail to trigger NMD because the
termination event would provide normal termination interactions between eRF3 and the
poly(A) binding protein (PABP in humans, Pab1 in yeast; Amrani et al., 2004; BehmAnsmant et al., 2007; Eberle et al., 2008; Ivanov et al., 2008; Singh et al., 2008).
When a PTC is located near the 3’ end of the β-globin mRNA, allowing the
mRNA to escape NMD, a C-terminally truncated polypeptide is produced that acts in a
dominant-negative manner and results in a severe form of heterozygous symptomatic βthalassemia (Thein et al., 1990, Hall and Thein 1994). In contrast, PTCs that are located
more 5’ in the β-globin mRNA are able to trigger NMD which limits the build-up of
toxic C-terminally truncated polypeptides. The result is an asymptomatic form of
heterozygous β-thalassemia (Nagy and Maquat, 1998; Thermann et al., 1998). However,
there is one particular set of mutations in the β-globin mRNA that do not fit this pattern.
Nonsense mutations in the first exon of the β-globin mRNA, which should activate
NMD, do not trigger degradation of the mRNA by NMD and do not result in
27
symptomatic β-thalassemia (Inacio et al., 2004; Romao et al., 2000). In 2011 Neu-Yilik
and colleagues solved this mystery by showing that PTCs in the first exon do not trigger
NMD because the ribosome is able to reinitiate translation in exon 2 at Met55. This
Met55 is the only in-frame downstream start codon and is located within an optimal
initiation context. Importantly, they showed that mutation of the downstream Met55 start
codon restores the NMD-sensitivity of the mRNA when a PTC is present upstream.
Additionally, PTCs that are located downstream of this Met55 do trigger NMD (NeuYilik et al., 2011). Together, these results confirm translation reinitiation as a method of
protecting mRNAs from degradation by NMD, but further examples have not yet been
demonstrated.
4.4 Trans-acting factor Pub1
Two examples of wild-type mRNAs with NMD targeting signals that are not
destabilized by NMD are the YAP1 and GCN4 mRNAs in S. cerevisiae. These two
mRNAs contain translated uORFs but show no significant difference in mRNA
accumulation or half-life (Michel et al., 2014; Ruiz-Echevarria et al., 1998; Vilela et al.,
1998; Chapter 4). A study by Ruiz-Echevarria and Peltz in 2000 identified the transacting factor poly(U)-binding protein, Pub1, as the mechanism responsible for the
protection of these two mRNAs from NMD. They determined that Pub1 bound to
stabilizer elements (STEs) located in the 5’ leader region of the mRNA. When Pub1 was
removed from the cells (pub1Δ strains) the mRNAs were significantly destabilized and,
importantly, this destabilization was dependent on NMD factor Upf1 (Ruiz-Echevarria
28
and Peltz, 2000). However, this data was not reproducible in a follow-up study
(unpublished data, Chapter 4).
5.0 Perspective and Discussion
Although originally discovered as a surveillance mechanism, further studies of
NMD provide evidence that this mechanism of mRNA turnover plays a significant role in
regulating the fragile and intricate process of eukaryotic gene regulation. When gene
regulation is in check and expression is occurring normally then organisms achieve
cellular homeostasis which leads to overall health. However, when gene regulation goes
awry it most often results in cellular malfunction and the development of disease. The
NMD pathway has been shown to play a role in the development of a significant number
of diseases. Interestingly, NMD has been shown to play both a protective and
aggravating role in the development of disease. If the C-terminally truncated proteins
produced from an mRNA with a PTC would be toxic to the cell then NMD serves a
protecting role. However, if the truncated protein would still retain enough activity to be
at least partially functional, then continually degrading the mRNA leads to a severe
deficiency of the protein. Because inherited PTCs play a role in the development of a
significant number of genetically related diseases and cancers researchers are actively
pursuing ways of modulating the cellular machinery in order to suppress or readthrough
the nonsense codon. Because PTCs can arise through a variety of mechanism (point
mutations, insertions or deletions, errors in alternative splicing) the approach to nonsense
suppression is not likely to be universal for all PTCs. Accordingly, it is much easier to
29
approach nonsense suppression of PTCs that have arisen due to a point mutation than it is
to approach the suppression of PTCs that have arisen in a way that would cause a shift in
the downstream reading frame. The latter of these also poses the complication
introducing multiple downstream PTCs due to the shift in reading frame.
It is important to point out that we currently do not have a thorough understanding
of the molecular mechanisms of NMD. Notably, it remains elusive as to exactly how a
cell is able to distinguish a normal termination event from a premature termination event.
Several models have been proposed but so far none of them are able to fully explain all of
the observed results of the summation of NMD studies. Considering the diversity of
results in studies aiming to elucidate the mechanism behind PTC versus normal
termination codon (NTC) differentiation it is reasonable to hypothesize that this
mechanism may be cell/tissue-specific. Indeed the faux 3’ UTR model seems to provide
the most comprehensive model and is supported by a significant number of studies and
observations.
Additionally, only recently have we begun to appreciate the full breadth of wildtype gene regulation by NMD. Genome-wide studies in various organisms all provide
evidence that NMD plays a significant role outside of mRNA surveillance. As the
studies of wild-type gene regulation by NMD continue to expand our knowledge we are
likely to gain a better understanding of additionally NMD-targeting mechanisms as well
as NMD-protection mechanisms. These studies are important as they may provide
valuable insight into understanding the underlying mechanisms of NMD.
30
Understanding the intricate molecular mechanisms of NMD will provide a more
solid foundation for the treatment of NMD-related diseases. Therapies can be developed
with greater efficiency and, with a better understanding of the overall role of NMD in the
cell, we can minimize side-effects and risks associated with disease therapy.
31
References
Aigner, S., Lingner, J., Goodrich, K.J., Grosshans, C.A., Shevchenko, A., Mann, M., and
Cech, T.R. (2000). Euplotes telomerase contains an La motif protein produced by
apparent translational frameshifting. EMBO J 19, 6230-6239.
Akimitsu, N. (2008). Messenger RNA surveillance systems monitoring proper translation
termination. J Biochem 143, 1-8.
Altamura, N., Groudinsky, O., Dujardin, G., and Slonimski, P.P. (1992). NAM7 nuclear
gene encodes a novel member of a family of helicases with a Zn-ligand motif and is
involved in mitochondrial functions in Saccharomyces cerevisiae. J Mol Biol 224, 575587.
Amrani, N., Dong, S., He, F., Ganesan, R., Ghosh, S., Kervestin, S., Li, C., Mangus,
D.A., Spatrick, P., and Jacobson, A. (2006). Aberrant termination triggers nonsensemediated mRNA decay. Biochem Soc Trans 34, 39-42.
Amrani, N., Ganesan, R., Kervestin, S., Mangus, D.A., Ghosh, S., and Jacobson, A.
(2004). A faux 3'-UTR promotes aberrant termination and triggers nonsense-mediated
mRNA decay. Nature 432, 112-118.
Applequist, S.E., Selg, M., Raman, C., and Jack, H.M. (1997). Cloning and
characterization of HUPF1, a human homolog of the Saccharomyces cerevisiae nonsense
mRNA-reducing UPF1 protein. Nucleic Acids Res 25, 814-821.
Arribere, J.A., and Gilbert, W.V. (2013). Roles for transcript leaders in translation and
mRNA decay revealed by transcript leader sequencing. Genome Res 23, 977-987.
Asakura, T., Sasaki, T., Nagano, F., Satoh, A., Obaishi, H., Nishioka, H., Imamura, H.,
Hotta, K., Tanaka, K., Nakanishi, H., et al. (1998). Isolation and characterization of a
novel actin filament-binding protein from Saccharomyces cerevisiae. Oncogene 16, 121130.
Atkin, A.L., Schenkman, L.R., Eastham, M., Dahlseid, J.N., Lelivelt, M.J., and
Culbertson, M.R. (1997). Relationship between yeast polyribosomes and Upf proteins
required for nonsense mRNA decay. J Biol Chem 272, 22163-22172.
Baker, K.E., and Parker, R. (2004). Nonsense-mediated mRNA decay: terminating
erroneous gene expression. Curr Opin Cell Biol 16, 293-299.
Baranov, P.V., Gesteland, R.F., and Atkins, J.F. (2002). Release factor 2 frameshifting
sites in different bacteria. EMBO Rep 3, 373-377.
Barbosa, C., Peixeiro, I., and Romao, L. (2013). Gene expression regulation by upstream
open reading frames and human disease. PLoS Genet 9, e1003529.
32
Baril, M., Dulude, D., Gendron, K., Lemay, G., and Brakier-Gingras, L. (2003).
Efficiency of a programmed -1 ribosomal frameshift in the different subtypes of the
human immunodeficiency virus type 1 group M. RNA 9, 1246-1253.
Bedwell, D.M., Kaenjak, A., Benos, D.J., Bebok, Z., Bubien, J.K., Hong, J., Tousson, A.,
Clancy, J.P., and Sorscher, E.J. (1997). Suppression of a CFTR premature stop mutation
in a bronchial epithelial cell line. Nat Med 3, 1280-1284.
Behm-Ansmant, I., Gatfield, D., Rehwinkel, J., Hilgers, V., and Izaurralde, E. (2007). A
conserved role for cytoplasmic poly(A)-binding protein 1 (PABPC1) in nonsensemediated mRNA decay. EMBO J 26, 1591-1601.
Beilharz, T.H., and Preiss, T. (2007). Widespread use of poly(A) tail length control to
accentuate expression of the yeast transcriptome. RNA 13, 982-997.
Bertram, G., Innes, S., Minella, O., Richardson, J., and Stansfield, I. (2001). Endless
possibilities: translation termination and stop codon recognition. Microbiology 147, 255269.
Bhattacharya, A., Czaplinski, K., Trifillis, P., He, F., Jacobson, A., and Peltz, S.W.
(2000). Characterization of the biochemical properties of the human Upf1 gene product
that is involved in nonsense-mediated mRNA decay. RNA 6, 1226-1235.
Bhuvanagiri, M., Schlitter, A.M., Hentze, M.W., and Kulozik, A.E. (2010). NMD: RNA
biology meets human genetic medicine. Biochem J 430, 365-377.
Blinkowa, A.L., and Walker, J.R. (1990). Programmed ribosomal frameshifting generates
the Escherichia coli DNA polymerase III gamma subunit from within the tau subunit
reading frame. Nucleic Acids Res 18, 1725-1729.
Bonetti, B., Fu, L., Moon, J., and Bedwell, D.M. (1995). The efficiency of translation
termination is determined by a synergistic interplay between upstream and downstream
sequences in Saccharomyces cerevisiae. J Mol Biol 251, 334-345.
Breitwieser, W., Price, C., and Schuster, T. (1993). Identification of a gene encoding a
novel zinc finger protein in Saccharomyces cerevisiae. Yeast 9, 551-556.
Brierley, I., and Dos Ramos, F.J. (2006). Programmed ribosomal frameshifting in HIV-1
and the SARS-CoV. Virus Res 119, 29-42.
Brogna, S., and Wen, J. (2009). Nonsense-mediated mRNA decay (NMD) mechanisms.
Nat Struct Mol Biol 16, 107-113.
Cali, B.M., Kuchma, S.L., Latham, J., and Anderson, P. (1999). smg-7 is required for
mRNA surveillance in Caenorhabditis elegans. Genetics 151, 605-616.
33
Calvo, S.E., Pagliarini, D.J., and Mootha, V.K. (2009). Upstream open reading frames
cause widespread reduction of protein expression and are polymorphic among humans.
Proc Natl Acad Sci U S A 106, 7507-7512.
Cao, A., and Galanello, R. (2010). Beta-thalassemia. Genet Med 12, 61-76.
Chamieh, H., Ballut, L., Bonneau, F., and Le Hir, H. (2008). NMD factors UPF2 and
UPF3 bridge UPF1 to the exon junction complex and stimulate its RNA helicase activity.
Nat Struct Mol Biol 15, 85-93.
Chang, Y.F., Imam, J.S., and Wilkinson, M.F. (2007). The nonsense-mediated decay
RNA surveillance pathway. Annu Rev Biochem 76, 51-74.
Cigan, A.M., Pabich, E.K., and Donahue, T.F. (1988). Mutational analysis of the HIS4
translational initiator region in Saccharomyces cerevisiae. Mol Cell Biol 8, 2964-2975.
Coller, J., and Parker, R. (2004). Eukaryotic mRNA decapping. Annu Rev Biochem 73,
861-890.
Cuccurese, M., Russo, G., Russo, A., and Pietropaolo, C. (2005). Alternative splicing and
nonsense-mediated mRNA decay regulate mammalian ribosomal gene expression.
Nucleic Acids Res 33, 5965-5977.
Cui, Y., Hagan, K.W., Zhang, S., and Peltz, S.W. (1995). Identification and
characterization of genes that are required for the accelerated degradation of mRNAs
containing a premature translational termination codon. Genes Dev 9, 423-436.
Culbertson, M.R. (1999). RNA surveillance. Unforeseen consequences for gene
expression, inherited genetic disorders and cancer. Trends Genet 15, 74-80.
Czaplinski, K., Ruiz-Echevarria, M.J., Gonzalez, C.I., and Peltz, S.W. (1999). Should we
kill the messenger? The role of the surveillance complex in translation termination and
mRNA turnover. Bioessays 21, 685-696.
Czaplinski, K., Ruiz-Echevarria, M.J., Paushkin, S.V., Han, X., Weng, Y., Perlick, H.A.,
Dietz, H.C., Ter-Avanesyan, M.D., and Peltz, S.W. (1998). The surveillance complex
interacts with the translation release factors to enhance termination and degrade aberrant
mRNAs. Genes Dev 12, 1665-1677.
Czaplinski, K., Weng, Y., Hagan, K.W., and Peltz, S.W. (1995). Purification and
characterization of the Upf1 protein: a factor involved in translation and mRNA
degradation. RNA 1, 610-623.
Decker, C.J., and Parker, R. (1993). A turnover pathway for both stable and unstable
mRNAs in yeast: evidence for a requirement for deadenylation. Genes Dev 7, 1632-1643.
34
Denning, G., Jamieson, L., Maquat, L.E., Thompson, E.A., and Fields, A.P. (2001).
Cloning of a novel phosphatidylinositol kinase-related kinase: characterization of the
human SMG-1 RNA surveillance protein. J Biol Chem 276, 22709-22714.
DePristo, M.A., Weinreich, D.M., and Hartl, D.L. (2005). Missense meanderings in
sequence space: a biophysical view of protein evolution. Nat Rev Genet 6, 678-687.
Dietz, H.C. (1993). Marfan Syndrome. In GeneReviews(R), R.A. Pagon, M.P. Adam,
H.H. Ardinger, S.E. Wallace, A. Amemiya, L.J.H. Bean, T.D. Bird, C.T. Fong, H.C.
Mefford, R.J.H. Smith, and K. Stephens, eds. (Seattle, WA).
Dietz, H.C., McIntosh, I., Sakai, L.Y., Corson, G.M., Chalberg, S.C., Pyeritz, R.E., and
Francomano, C.A. (1993). Four novel FBN1 mutations: significance for mutant transcript
level and EGF-like domain calcium binding in the pathogenesis of Marfan syndrome.
Genomics 17, 468-475.
Dinman, J.D. (2012). Control of gene expression by translational recoding. Adv Protein
Chem Struct Biol 86, 129-149.
Dinman, J.D., Icho, T., and Wickner, R.B. (1991). A -1 ribosomal frameshift in a doublestranded RNA virus of yeast forms a gag-pol fusion protein. Proc Natl Acad Sci U S A
88, 174-178.
Dulude, D., Baril, M., and Brakier-Gingras, L. (2002). Characterization of the frameshift
stimulatory signal controlling a programmed -1 ribosomal frameshift in the human
immunodeficiency virus type 1. Nucleic Acids Res 30, 5094-5102.
Eberle, A.B., Stalder, L., Mathys, H., Orozco, R.Z., and Muhlemann, O. (2008).
Posttranscriptional gene regulation by spatial rearrangement of the 3' untranslated region.
PLoS Biol 6, e92.
Fearon, K., McClendon, V., Bonetti, B., and Bedwell, D.M. (1994). Premature translation
termination mutations are efficiently suppressed in a highly conserved region of yeast
Ste6p, a member of the ATP-binding cassette (ABC) transporter family. J Biol Chem
269, 17802-17808.
Finkel, R.S. (2010). Read-through strategies for suppression of nonsense mutations in
Duchenne/ Becker muscular dystrophy: aminoglycosides and ataluren (PTC124). J Child
Neurol 25, 1158-1164.
Frischmeyer, P.A., and Dietz, H.C. (1999). Nonsense-mediated mRNA decay in health
and disease. Hum Mol Genet 8, 1893-1900.
Gaba, A., Jacobson, A., and Sachs, M.S. (2005). Ribosome occupancy of the yeast CPA1
upstream open reading frame termination codon modulates nonsense-mediated mRNA
decay. Mol Cell 20, 449-460.
35
Gallie, D.R. (1991). The cap and poly(A) tail function synergistically to regulate mRNA
translational efficiency. Genes Dev 5, 2108-2116.
Gatfield, D., Unterholzner, L., Ciccarelli, F.D., Bork, P., and Izaurralde, E. (2003).
Nonsense-mediated mRNA decay in Drosophila: at the intersection of the yeast and
mammalian pathways. EMBO J 22, 3960-3970.
Gozalbo, D., and Hohmann, S. (1990). Nonsense suppressors partially revert the decrease
of the mRNA level of a nonsense mutant allele in yeast. Curr Genet 17, 77-79.
Graber, J.H., Cantor, C.R., Mohr, S.C., and Smith, T.F. (1999). Genomic detection of
new yeast pre-mRNA 3'-end-processing signals. Nucleic Acids Res 27, 888-894.
Graschopf, A., Stadler, J.A., Hoellerer, M.K., Eder, S., Sieghardt, M., Kohlwein, S.D.,
and Schweyen, R.J. (2001). The yeast plasma membrane protein Alr1 controls Mg2+
homeostasis and is subject to Mg2+-dependent control of its synthesis and degradation. J
Biol Chem 276, 16216-16222.
Grimson, A., O'Connor, S., Newman, C.L., and Anderson, P. (2004). SMG-1 is a
phosphatidylinositol kinase-related protein kinase required for nonsense-mediated mRNA
Decay in Caenorhabditis elegans. Mol Cell Biol 24, 7483-7490.
Guan, Q., Zheng, W., Tang, S., Liu, X., Zinkel, R.A., Tsui, K.W., Yandell, B.S., and
Culbertson, M.R. (2006). Impact of nonsense-mediated mRNA decay on the global
expression profile of budding yeast. PLoS Genet 2, e203.
Gunn, G., Dai, Y., Du, M., Belakhov, V., Kandasamy, J., Schoeb, T.R., Baasov, T.,
Bedwell, D.M., and Keeling, K.M. (2014). Long-term nonsense suppression therapy
moderates MPS I-H disease progression. Mol Genet Metab 111, 374-381.
Hall, G.W., and Thein, S. (1994). Nonsense codon mutations in the terminal exon of the
beta-globin gene are not associated with a reduction in beta-mRNA accumulation: a
mechanism for the phenotype of dominant beta-thalassemia. Blood 83, 2031-2037.
Hamilton, R., Watanabe, C.K., and de Boer, H.A. (1987). Compilation and comparison of
the sequence context around the AUG startcodons in Saccharomyces cerevisiae mRNAs.
Nucleic Acids Res 15, 3581-3593.
Hammell, A.B., Taylor, R.C., Peltz, S.W., and Dinman, J.D. (1999). Identification of
putative programmed -1 ribosomal frameshift signals in large DNA databases. Genome
Res 9, 417-427.
Hansen, K.D., Lareau, L.F., Blanchette, M., Green, R.E., Meng, Q., Rehwinkel, J.,
Gallusser, F.L., Izaurralde, E., Rio, D.C., Dudoit, S., et al. (2009). Genome-wide
identification of alternative splice forms down-regulated by nonsense-mediated mRNA
decay in Drosophila. PLoS Genet 5, e1000525.
36
Hansen, T.M., Reihani, S.N., Oddershede, L.B., and Sorensen, M.A. (2007). Correlation
between mechanical strength of messenger RNA pseudoknots and ribosomal
frameshifting. Proc Natl Acad Sci U S A 104, 5830-5835.
He, F., Brown, A.H., and Jacobson, A. (1997). Upf1p, Nmd2p, and Upf3p are interacting
components of the yeast nonsense-mediated mRNA decay pathway. Mol Cell Biol 17,
1580-1594.
He, F., and Jacobson, A. (1995). Identification of a novel component of the nonsensemediated mRNA decay pathway by use of an interacting protein screen. Genes Dev 9,
437-454.
He, F., Li, X., Spatrick, P., Casillo, R., Dong, S., and Jacobson, A. (2003). Genome-wide
analysis of mRNAs regulated by the nonsense-mediated and 5' to 3' mRNA decay
pathways in yeast. Mol Cell 12, 1439-1452.
He, F., Peltz, S.W., Donahue, J.L., Rosbash, M., and Jacobson, A. (1993). Stabilization
and ribosome association of unspliced pre-mRNAs in a yeast upf1- mutant. Proc Natl
Acad Sci U S A 90, 7034-7038.
Hentze, M.W., and Kulozik, A.E. (1999). A perfect message: RNA surveillance and
nonsense-mediated decay. Cell 96, 307-310.
Herrick, D., Parker, R., and Jacobson, A. (1990). Identification and comparison of stable
and unstable mRNAs in Saccharomyces cerevisiae. Mol Cell Biol 10, 2269-2284.
Hilleren, P., and Parker, R. (1999). Mechanisms of mRNA surveillance in eukaryotes.
Annu Rev Genet 33, 229-260.
Hodgkin, J., Papp, A., Pulak, R., Ambros, V., and Anderson, P. (1989). A new kind of
informational suppression in the nematode Caenorhabditis elegans. Genetics 123, 301313.
Hsu, C.L., and Stevens, A. (1993). Yeast cells lacking 5'-->3' exoribonuclease 1 contain
mRNA species that are poly(A) deficient and partially lack the 5' cap structure. Mol Cell
Biol 13, 4826-4835.
Hurt, J.A., Robertson, A.D., and Burge, C.B. (2013). Global analyses of UPF1 binding
and function reveal expanded scope of nonsense-mediated mRNA decay. Genome Res
23, 1636-1650.
Inacio, A., Silva, A.L., Pinto, J., Ji, X., Morgado, A., Almeida, F., Faustino, P., Lavinha,
J., Liebhaber, S.A., and Romao, L. (2004). Nonsense mutations in close proximity to the
initiation codon fail to trigger full nonsense-mediated mRNA decay. J Biol Chem 279,
32170-32180.
Isken, O., and Maquat, L.E. (2008). The multiple lives of NMD factors: balancing roles
in gene and genome regulation. Nat Rev Genet 9, 699-712.
37
Ivanov, I.P., Matsufuji, S., Murakami, Y., Gesteland, R.F., and Atkins, J.F. (2000).
Conservation of polyamine regulation by translational frameshifting from yeast to
mammals. EMBO J 19, 1907-1917.
Ivanov, P.V., Gehring, N.H., Kunz, J.B., Hentze, M.W., and Kulozik, A.E. (2008).
Interactions between UPF1, eRFs, PABP and the exon junction complex suggest an
integrated model for mammalian NMD pathways. EMBO J 27, 736-747.
Jacks, T., and Varmus, H.E. (1985). Expression of the Rous sarcoma virus pol gene by
ribosomal frameshifting. Science 230, 1237-1242.
Jaillon, O., Bouhouche, K., Gout, J.F., Aury, J.M., Noel, B., Saudemont, B., Nowacki,
M., Serrano, V., Porcel, B.M., Segurens, B., et al. (2008). Translational control of intron
splicing in eukaryotes. Nature 451, 359-362.
Jeong, H.J., Kim, Y.J., Kim, S.H., Kim, Y.H., Lee, I.J., Kim, Y.K., and Shin, J.S. (2011).
Nonsense-mediated mRNA decay factors, UPF1 and UPF3, contribute to plant defense.
Plant Cell Physiol 52, 2147-2156.
Johansson, M.J., He, F., Spatrick, P., Li, C., and Jacobson, A. (2007). Association of
yeast Upf1p with direct substrates of the NMD pathway. Proc Natl Acad Sci U S A 104,
20872-20877.
Johansson, M.J., and Jacobson, A. (2010). Nonsense-mediated mRNA decay maintains
translational fidelity by limiting magnesium uptake. Genes Dev 24, 1491-1495.
Kadlec, J., Guilligay, D., Ravelli, R.B., and Cusack, S. (2006). Crystal structure of the
UPF2-interacting domain of nonsense-mediated mRNA decay factor UPF1. RNA 12,
1817-1824.
Kalyna, M., Simpson, C.G., Syed, N.H., Lewandowska, D., Marquez, Y., Kusenda, B.,
Marshall, J., Fuller, J., Cardle, L., McNicol, J., et al. (2012). Alternative splicing and
nonsense-mediated decay modulate expression of important regulatory genes in
Arabidopsis. Nucleic Acids Res 40, 2454-2469.
Kawashima, T., Douglass, S., Gabunilas, J., Pellegrini, M., and Chanfreau, G.F. (2014).
Widespread use of non-productive alternative splice sites in Saccharomyces cerevisiae.
PLoS Genet 10, e1004249.
Kebaara, B.W., and Atkin, A.L. (2009). Long 3'-UTRs target wild-type mRNAs for
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Nucleic Acids Res 37,
2771-2778.
Keeling, K.M., Lanier, J., Du, M., Salas-Marco, J., Gao, L., Kaenjak-Angeletti, A., and
Bedwell, D.M. (2004). Leaky termination at premature stop codons antagonizes
nonsense-mediated mRNA decay in S. cerevisiae. RNA 10, 691-703.
38
Kerr, T.P., Duward, A., Hodgson, S.V., Hughes, E., and Robb, S.A. (2001).
Hyperkalaemic cardiac arrest in a manifesting carrier of Duchenne muscular dystrophy
following general anaesthesia. Eur J Pediatr 160, 579-580.
Kertesz, S., Kerenyi, Z., Merai, Z., Bartos, I., Palfy, T., Barta, E., and Silhavy, D. (2006).
Both introns and long 3'-UTRs operate as cis-acting elements to trigger nonsensemediated decay in plants. Nucleic Acids Res 34, 6147-6157.
Khajavi, M., Inoue, K., and Lupski, J.R. (2006). Nonsense-mediated mRNA decay
modulates clinical outcome of genetic disease. Eur J Hum Genet 14, 1074-1081.
Kisselev, L., Ehrenberg, M., and Frolova, L. (2003). Termination of translation: interplay
of mRNA, rRNAs and release factors? EMBO J 22, 175-182.
Kochetov, A.V., Syrnik, O.A., Rogozin, I.B., Glazko, G.V., Komarova, M.L., and
Shumnyi, V.K. (2002). [Context organization of mRNA 5'-untranslated regions of higher
plants]. Mol Biol (Mosk) 36, 649-656.
Kurihara, Y., Matsui, A., Hanada, K., Kawashima, M., Ishida, J., Morosawa, T., Tanaka,
M., Kaminuma, E., Mochizuki, Y., Matsushima, A., et al. (2009). Genome-wide
suppression of aberrant mRNA-like noncoding RNAs by NMD in Arabidopsis. Proc Natl
Acad Sci U S A 106, 2453-2458.
Kuzmiak, H.A., and Maquat, L.E. (2006). Applying nonsense-mediated mRNA decay
research to the clinic: progress and challenges. Trends Mol Med 12, 306-316.
Le Hir, H., Gatfield, D., Izaurralde, E., and Moore, M.J. (2001). The exon-exon junction
complex provides a binding platform for factors involved in mRNA export and nonsensemediated mRNA decay. EMBO J 20, 4987-4997.
Lee, B.S., and Culbertson, M.R. (1995). Identification of an additional gene required for
eukaryotic nonsense mRNA turnover. Proc Natl Acad Sci U S A 92, 10354-10358.
Leeds, P., Peltz, S.W., Jacobson, A., and Culbertson, M.R. (1991). The product of the
yeast UPF1 gene is required for rapid turnover of mRNAs containing a premature
translational termination codon. Genes Dev 5, 2303-2314.
Leeds, P., Wood, J.M., Lee, B.S., and Culbertson, M.R. (1992). Gene products that
promote mRNA turnover in Saccharomyces cerevisiae. Mol Cell Biol 12, 2165-2177.
Leger, M., Dulude, D., Steinberg, S.V., and Brakier-Gingras, L. (2007). The three
transfer RNAs occupying the A, P and E sites on the ribosome are involved in viral
programmed -1 ribosomal frameshift. Nucleic Acids Res 35, 5581-5592.
Lejeune, F., Cavaloc, Y., and Stevenin, J. (2001). Alternative splicing of intron 3 of the
serine/arginine-rich protein 9G8 gene. Identification of flanking exonic splicing
enhancers and involvement of 9G8 as a trans-acting factor. J Biol Chem 276, 7850-7858.
39
Lelivelt, M.J., and Culbertson, M.R. (1999). Yeast Upf proteins required for RNA
surveillance affect global expression of the yeast transcriptome. Mol Cell Biol 19, 67106719.
Lewis, B.P., Green, R.E., and Brenner, S.E. (2003). Evidence for the widespread
coupling of alternative splicing and nonsense-mediated mRNA decay in humans. Proc
Natl Acad Sci U S A 100, 189-192.
Longman, D., Hug, N., Keith, M., Anastasaki, C., Patton, E.E., Grimes, G., and Caceres,
J.F. (2013). DHX34 and NBAS form part of an autoregulatory NMD circuit that regulates
endogenous RNA targets in human cells, zebrafish and Caenorhabditis elegans. Nucleic
Acids Res 41, 8319-8331.
Lykke-Andersen, J. (2002). Identification of a human decapping complex associated with
hUpf proteins in nonsense-mediated decay. Mol Cell Biol 22, 8114-8121.
Lykke-Andersen, J., and Bennett, E.J. (2014). Protecting the proteome: Eukaryotic
cotranslational quality control pathways. J Cell Biol 204, 467-476.
Lykke-Andersen, J., Shu, M.D., and Steitz, J.A. (2000). Human Upf proteins target an
mRNA for nonsense-mediated decay when bound downstream of a termination codon.
Cell 103, 1121-1131.
Lykke-Andersen, J., Shu, M.D., and Steitz, J.A. (2001). Communication of the position
of exon-exon junctions to the mRNA surveillance machinery by the protein RNPS1.
Science 293, 1836-1839.
MacDiarmid, C.W., and Gardner, R.C. (1998). Overexpression of the Saccharomyces
cerevisiae magnesium transport system confers resistance to aluminum ion. J Biol Chem
273, 1727-1732.
Maderazo, A.B., He, F., Mangus, D.A., and Jacobson, A. (2000). Upf1p control of
nonsense mRNA translation is regulated by Nmd2p and Upf3p. Mol Cell Biol 20, 45914603.
Maquat, L.E., and Carmichael, G.G. (2001). Quality control of mRNA function. Cell 104,
173-176.
Maquat, L.E., and Serin, G. (2001). Nonsense-mediated mRNA decay: insights into
mechanism from the cellular abundance of human Upf1, Upf2, Upf3, and Upf3X
proteins. Cold Spring Harb Symp Quant Biol 66, 313-320.
McGlincy, N.J., and Smith, C.W. (2008). Alternative splicing resulting in nonsensemediated mRNA decay: what is the meaning of nonsense? Trends Biochem Sci 33, 385393.
Meaux, S., van Hoof, A., and Baker, K.E. (2008). Nonsense-mediated mRNA decay in
yeast does not require PAB1 or a poly(A) tail. Mol Cell 29, 134-140.
40
Mendell, J.T., ap Rhys, C.M., and Dietz, H.C. (2002). Separable roles for rent1/hUpf1 in
altered splicing and decay of nonsense transcripts. Science 298, 419-422.
Mendell, J.T., Sharifi, N.A., Meyers, J.L., Martinez-Murillo, F., and Dietz, H.C. (2004).
Nonsense surveillance regulates expression of diverse classes of mammalian transcripts
and mutes genomic noise. Nat Genet 36, 1073-1078.
Michel, A.M., Fox, G., A, M.K., De Bo, C., O'Connor, P.B., Heaphy, S.M., Mullan, J.P.,
Donohue, C.A., Higgins, D.G., and Baranov, P.V. (2014). GWIPS-viz: development of a
ribo-seq genome browser. Nucleic Acids Res 42, D859-864.
Mitrovich, Q.M., and Anderson, P. (2000). Unproductively spliced ribosomal protein
mRNAs are natural targets of mRNA surveillance in C. elegans. Genes Dev 14, 21732184.
Mitrovich, Q.M., and Anderson, P. (2005). mRNA surveillance of expressed
pseudogenes in C. elegans. Curr Biol 15, 963-967.
Miyasaka, H. (1999). The positive relationship between codon usage bias and translation
initiation AUG context in Saccharomyces cerevisiae. Yeast 15, 633-637.
Morris, D.K., and Lundblad, V. (1997). Programmed translational frameshifting in a gene
required for yeast telomere replication. Curr Biol 7, 969-976.
Muhlemann, O., Eberle, A.B., Stalder, L., and Zamudio Orozco, R. (2008). Recognition
and elimination of nonsense mRNA. Biochim Biophys Acta 1779, 538-549.
Muhlrad, D., Decker, C.J., and Parker, R. (1994). Deadenylation of the unstable mRNA
encoded by the yeast MFA2 gene leads to decapping followed by 5'-->3' digestion of the
transcript. Genes Dev 8, 855-866.
Muhlrad, D., Decker, C.J., and Parker, R. (1995). Turnover mechanisms of the stable
yeast PGK1 mRNA. Mol Cell Biol 15, 2145-2156.
Muhlrad, D., and Parker, R. (1999). Aberrant mRNAs with extended 3' UTRs are
substrates for rapid degradation by mRNA surveillance. RNA 5, 1299-1307.
Nagalakshmi, U., Wang, Z., Waern, K., Shou, C., Raha, D., Gerstein, M., and Snyder, M.
(2008). The transcriptional landscape of the yeast genome defined by RNA sequencing.
Science 320, 1344-1349.
Nagy, E., and Maquat, L.E. (1998). A rule for termination-codon position within introncontaining genes: when nonsense affects RNA abundance. Trends Biochem Sci 23, 198199.
Namy, O., Duchateau-Nguyen, G., and Rousset, J.P. (2002). Translational readthrough of
the PDE2 stop codon modulates cAMP levels in Saccharomyces cerevisiae. Mol
Microbiol 43, 641-652.
41
Namy, O., Hatin, I., and Rousset, J.P. (2001). Impact of the six nucleotides downstream
of the stop codon on translation termination. EMBO Rep 2, 787-793.
Namy, O., Moran, S.J., Stuart, D.I., Gilbert, R.J., and Brierley, I. (2006). A mechanical
explanation of RNA pseudoknot function in programmed ribosomal frameshifting.
Nature 441, 244-247.
Namy, O., Rousset, J.P., Napthine, S., and Brierley, I. (2004). Reprogrammed genetic
decoding in cellular gene expression. Mol Cell 13, 157-168.
Neu-Yilik, G., Amthor, B., Gehring, N.H., Bahri, S., Paidassi, H., Hentze, M.W., and
Kulozik, A.E. (2011). Mechanism of escape from nonsense-mediated mRNA decay of
human beta-globin transcripts with nonsense mutations in the first exon. RNA 17, 843854.
Ni, J.Z., Grate, L., Donohue, J.P., Preston, C., Nobida, N., O'Brien, G., Shiue, L., Clark,
T.A., Blume, J.E., and Ares, M., Jr. (2007). Ultraconserved elements are associated with
homeostatic control of splicing regulators by alternative splicing and nonsense-mediated
decay. Genes Dev 21, 708-718.
Nicholson, P., Yepiskoposyan, H., Metze, S., Zamudio Orozco, R., Kleinschmidt, N., and
Muhlemann, O. (2010). Nonsense-mediated mRNA decay in human cells: mechanistic
insights, functions beyond quality control and the double-life of NMD factors. Cell Mol
Life Sci 67, 677-700.
Nyiko, T., Sonkoly, B., Merai, Z., Benkovics, A.H., and Silhavy, D. (2009). Plant
upstream ORFs can trigger nonsense-mediated mRNA decay in a size-dependent manner.
Plant Mol Biol 71, 367-378.
Obenoskey, J., Lane, D.R., Atkin, A.L., and Kebaara, B.W. (2014). Immunity of the
Saccharomyces cerevisiae SSY5 mRNA to nonsense-mediated mRNA decay. Front Mol
Biosci 1, 25.
Peccarelli, M., and Kebaara, B.W. (2014). Regulation of natural mRNAs by the
nonsense-mediated mRNA decay pathway. Eukaryot Cell 13, 1126-1135.
Peixeiro, I., Silva, A.L., and Romao, L. (2011). Control of human beta-globin mRNA
stability and its impact on beta-thalassemia phenotype. Haematologica 96, 905-913.
Peltz, S.W., Brown, A.H., and Jacobson, A. (1993). mRNA destabilization triggered by
premature translational termination depends on at least three cis-acting sequence
elements and one trans-acting factor. Genes Dev 7, 1737-1754.
Peltz, S.W., Donahue, J.L., and Jacobson, A. (1992). A mutation in the tRNA
nucleotidyltransferase gene promotes stabilization of mRNAs in Saccharomyces
cerevisiae. Mol Cell Biol 12, 5778-5784.
42
Peltz, S.W., Morsy, M., Welch, E.M., and Jacobson, A. (2013). Ataluren as an agent for
therapeutic nonsense suppression. Annu Rev Med 64, 407-425.
Pierard, A., Messenguy, F., Feller, A., and Hilger, F. (1979). Dual regulation of the
synthesis of the arginine pathway carbamoylphosphate synthase of Saccharomyces
cerevisiae by specific and general controls of amino acid biosynthesis. Mol Gen Genet
174, 163-171.
Plant, E.P., Wang, P., Jacobs, J.L., and Dinman, J.D. (2004). A programmed -1 ribosomal
frameshift signal can function as a cis-acting mRNA destabilizing element. Nucleic Acids
Res 32, 784-790.
Pulak, R., and Anderson, P. (1993). mRNA surveillance by the Caenorhabditis elegans
smg genes. Genes Dev 7, 1885-1897.
Ramani, A.K., Nelson, A.C., Kapranov, P., Bell, I., Gingeras, T.R., and Fraser, A.G.
(2009). High resolution transcriptome maps for wild-type and nonsense-mediated decaydefective Caenorhabditis elegans. Genome Biol 10, R101.
Rayson, S., Arciga-Reyes, L., Wootton, L., De Torres Zabala, M., Truman, W., Graham,
N., Grant, M., and Davies, B. (2012). A role for nonsense-mediated mRNA decay in
plants: pathogen responses are induced in Arabidopsis thaliana NMD mutants. PLoS One
7, e31917.
Rebbapragada, I., and Lykke-Andersen, J. (2009). Execution of nonsense-mediated
mRNA decay: what defines a substrate? Curr Opin Cell Biol 21, 394-402.
Rehwinkel, J., Letunic, I., Raes, J., Bork, P., and Izaurralde, E. (2005). Nonsensemediated mRNA decay factors act in concert to regulate common mRNA targets. RNA
11, 1530-1544.
Rogan, M.P., Stoltz, D.A., and Hornick, D.B. (2011). Cystic fibrosis transmembrane
conductance regulator intracellular processing, trafficking, and opportunities for
mutation-specific treatment. Chest 139, 1480-1490.
Romao, L., Inacio, A., Santos, S., Avila, M., Faustino, P., Pacheco, P., and Lavinha, J.
(2000). Nonsense mutations in the human beta-globin gene lead to unexpected levels of
cytoplasmic mRNA accumulation. Blood 96, 2895-2901.
Roque, S., Cerciat, M., Gaugue, I., Mora, L., Floch, A.G., de Zamaroczy, M., HeurgueHamard, V., and Kervestin, S. (2015). Interaction between the poly(A)-binding protein
Pab1 and the eukaryotic release factor eRF3 regulates translation termination but not
mRNA decay in Saccharomyces cerevisiae. RNA 21, 124-134.
Rospert, S., Rakwalska, M., and Dubaquie, Y. (2005). Polypeptide chain termination and
stop codon readthrough on eukaryotic ribosomes. Rev Physiol Biochem Pharmacol 155,
1-30.
43
Ruiz-Echevarria, M.J., Gonzalez, C.I., and Peltz, S.W. (1998). Identifying the right stop:
determining how the surveillance complex recognizes and degrades an aberrant mRNA.
EMBO J 17, 575-589.
Ruiz-Echevarria, M.J., and Peltz, S.W. (2000). The RNA binding protein Pub1 modulates
the stability of transcripts containing upstream open reading frames. Cell 101, 741-751.
Sato, K., Noda, Y., and Yoda, K. (2007). Pga1 is an essential component of
Glycosylphosphatidylinositol-mannosyltransferase II of Saccharomyces cerevisiae. Mol
Biol Cell 18, 3472-3485.
Schweingruber, C., Rufener, S.C., Zund, D., Yamashita, A., and Muhlemann, O. (2013).
Nonsense-mediated mRNA decay - mechanisms of substrate mRNA recognition and
degradation in mammalian cells. Biochim Biophys Acta 1829, 612-623.
Serin, G., Gersappe, A., Black, J.D., Aronoff, R., and Maquat, L.E. (2001). Identification
and characterization of human orthologues to Saccharomyces cerevisiae Upf2 protein and
Upf3 protein (Caenorhabditis elegans SMG-4). Mol Cell Biol 21, 209-223.
Sharp, P.M., and Li, W.H. (1987). The codon Adaptation Index--a measure of directional
synonymous codon usage bias, and its potential applications. Nucleic Acids Res 15,
1281-1295.
Shibuya, T., Tange, T.O., Sonenberg, N., and Moore, M.J. (2004). eIF4AIII binds spliced
mRNA in the exon junction complex and is essential for nonsense-mediated decay. Nat
Struct Mol Biol 11, 346-351.
Shirley, R.L., Lelivelt, M.J., Schenkman, L.R., Dahlseid, J.N., and Culbertson, M.R.
(1998). A factor required for nonsense-mediated mRNA decay in yeast is exported from
the nucleus to the cytoplasm by a nuclear export signal sequence. J Cell Sci 111 ( Pt 21),
3129-3143.
Shyu, A.B., Wilkinson, M.F., and van Hoof, A. (2008). Messenger RNA regulation: to
translate or to degrade. EMBO J 27, 471-481.
Silva, A.L., Ribeiro, P., Inacio, A., Liebhaber, S.A., and Romao, L. (2008). Proximity of
the poly(A)-binding protein to a premature termination codon inhibits mammalian
nonsense-mediated mRNA decay. RNA 14, 563-576.
Singh, G., Jakob, S., Kleedehn, M.G., and Lykke-Andersen, J. (2007). Communication
with the exon-junction complex and activation of nonsense-mediated decay by human
Upf proteins occur in the cytoplasm. Mol Cell 27, 780-792.
Singh, G., Rebbapragada, I., and Lykke-Andersen, J. (2008). A competition between
stimulators and antagonists of Upf complex recruitment governs human nonsensemediated mRNA decay. PLoS Biol 6, e111.
44
Sun, X., and Maquat, L.E. (2000). mRNA surveillance in mammalian cells: the
relationship between introns and translation termination. RNA 6, 1-8.
Sureau, A., Gattoni, R., Dooghe, Y., Stevenin, J., and Soret, J. (2001). SC35
autoregulates its expression by promoting splicing events that destabilize its mRNAs.
EMBO J 20, 1785-1796.
Takahashi, S., Araki, Y., Sakuno, T., and Katada, T. (2003). Interaction between Ski7p
and Upf1p is required for nonsense-mediated 3'-to-5' mRNA decay in yeast. EMBO J 22,
3951-3959.
Thein, S.L., Hesketh, C., Taylor, P., Temperley, I.J., Hutchinson, R.M., Old, J.M., Wood,
W.G., Clegg, J.B., and Weatherall, D.J. (1990a). Molecular basis for dominantly
inherited inclusion body beta-thalassemia. Proc Natl Acad Sci U S A 87, 3924-3928.
Thein, S.L., Winichagoon, P., Hesketh, C., Best, S., Fucharoen, S., Wasi, P., and
Weatherall, D.J. (1990b). The molecular basis of beta-thalassemia in Thailand:
application to prenatal diagnosis. Am J Hum Genet 47, 369-375.
Thermann, R., Neu-Yilik, G., Deters, A., Frede, U., Wehr, K., Hagemeier, C., Hentze,
M.W., and Kulozik, A.E. (1998). Binary specification of nonsense codons by splicing
and cytoplasmic translation. EMBO J 17, 3484-3494.
Tholstrup, J., Oddershede, L.B., and Sorensen, M.A. (2012). mRNA pseudoknot
structures can act as ribosomal roadblocks. Nucleic Acids Res 40, 303-313.
Vilela, C., Linz, B., Rodrigues-Pousada, C., and McCarthy, J.E. (1998). The yeast
transcription factor genes YAP1 and YAP2 are subject to differential control at the levels
of both translation and mRNA stability. Nucleic Acids Res 26, 1150-1159.
Weischenfeldt, J., Waage, J., Tian, G., Zhao, J., Damgaard, I., Jakobsen, J.S., Kristiansen,
K., Krogh, A., Wang, J., and Porse, B.T. (2012). Mammalian tissues defective in
nonsense-mediated mRNA decay display highly aberrant splicing patterns. Genome Biol
13, R35.
Welch, E.M., and Jacobson, A. (1999). An internal open reading frame triggers nonsensemediated decay of the yeast SPT10 mRNA. EMBO J 18, 6134-6145.
Weng, Y., Czaplinski, K., and Peltz, S.W. (1996a). Genetic and biochemical
characterization of mutations in the ATPase and helicase regions of the Upf1 protein.
Mol Cell Biol 16, 5477-5490.
Weng, Y., Czaplinski, K., and Peltz, S.W. (1996b). Identification and characterization of
mutations in the UPF1 gene that affect nonsense suppression and the formation of the
Upf protein complex but not mRNA turnover. Mol Cell Biol 16, 5491-5506.
45
Williams, I., Richardson, J., Starkey, A., and Stansfield, I. (2004). Genome-wide
prediction of stop codon readthrough during translation in the yeast Saccharomyces
cerevisiae. Nucleic Acids Res 32, 6605-6616.
Wilusz, C.J., Gao, M., Jones, C.L., Wilusz, J., and Peltz, S.W. (2001). Poly(A)-binding
proteins regulate both mRNA deadenylation and decapping in yeast cytoplasmic extracts.
RNA 7, 1416-1424.
Wollerton, M.C., Gooding, C., Wagner, E.J., Garcia-Blanco, M.A., and Smith, C.W.
(2004). Autoregulation of polypyrimidine tract binding protein by alternative splicing
leading to nonsense-mediated decay. Mol Cell 13, 91-100.
Yepiskoposyan, H., Aeschimann, F., Nilsson, D., Okoniewski, M., and Muhlemann, O.
(2011). Autoregulation of the nonsense-mediated mRNA decay pathway in human cells.
RNA 17, 2108-2118.
Yun, D.F., Laz, T.M., Clements, J.M., and Sherman, F. (1996). mRNA sequences
influencing translation and the selection of AUG initiator codons in the yeast
Saccharomyces cerevisiae. Mol Microbiol 19, 1225-1239.
Zhang, S., Welch, E.M., Hogan, K., Brown, A.H., Peltz, S.W., and Jacobson, A. (1997).
Polysome-associated mRNAs are substrates for the nonsense-mediated mRNA decay
pathway in Saccharomyces cerevisiae. RNA 3, 234-244.
Zuk, D., and Jacobson, A. (1998). A single amino acid substitution in yeast eIF-5A
results in mRNA stabilization. EMBO J 17, 2914-2925.
46
CHAPTER 2
Saccharomyces cerevisiae SSY5 mRNA is a wild-type mRNA with multiple NMDtargeting signals but is not degraded by NMD
47
Abstract
Nonsense-mediated mRNA decay is a translation-dependent surveillance
mechanism responsible for rapidly degrading mRNAs with premature termination codons
(PTCs). However, there is a significant portion of mRNAs that do not contain a PTC but
are substrates for the NMD pathway. The underlying mechanisms of how the cellular
machinery determines whether or not to degrade an mRNA via the NMD pathway are not
well understood. Here we present the case of the Saccharomyces cerevisiae SSY5
mRNA, which is a wild-type mRNA containing multiple NMD-targeting signals, but is
not degraded by the NMD pathway. We demonstrate that known mechanisms for the
protection of mRNAs from NMD do not apply to SSY5 mRNA as the mRNA is translated
and we do not find evidence of stop codon readthrough or translation reinitiation. Thus,
the protection of the SSY5 mRNA from NMD is likely due to a novel mechanism that
may provide further clues in unraveling the mystery of gene regulation and substrate
recognition by NMD.
48
Introduction
Nonsense-mediated mRNA decay (NMD) was historically identified as a posttranscriptional surveillance mechanism responsible for the rapid degradation of mRNAs
containing premature termination codons (PTCs; Gonzalez et al., 2001; Isken and
Maquat, 2008; Muhlemann et al., 2008). The identification of a termination event as
premature and the subsequent activation of the NMD machinery is critical for cellular
homeostasis as this process ensures that truncated proteins, which can otherwise have
dominant-negative deleterious effects, are not allowed to build up in the cell (Hentze and
Kulozik, 1999; Hilleren and Parker, 1999; Maquat and Serin, 2001). Additionally, a role
for NMD is implicated in as many as one third of inherited genetic diseases and cancers
(Frischmeyer and Dietz, 1999).
NMD requires the coordinated activity of three core factors: Upf1, Upf2 and
Upf3. Mutations or deletions in one or more of the genes encoding these factors
stabilizes NMD substrates (Cui et al., 1995; He et al., 1997; He and Jacobson, 1995; Lee
and Culbertson, 1995; Maderazo et al., 2000). Importantly, NMD is conserved from
yeast to humans and has been observed in all eukaryotes examined (Bedwell et al., 1997;
Hall and Thein, 1994; He et al., 2003; Hentze and Kulozik, 1999; Maquat and
Carmichael, 2001; Maquat and Serin, 2001; Mendell et al., 2004; Pulak and Anderson,
1993; Rehwinkel et al., 2005). However, the mechanism underlying the identification of
a termination event as premature and the subsequent rapid activation of the NMD
pathway remains largely obscure. Moreover, it has also been observed that NMD has an
impact on the accumulation of 3-10% of yeast mRNAs, many of which do not contain
49
PTCs (Culbertson, 1999; Guan et al., 2006; Lelivelt and Culbertson, 1999). Thus, we are
beginning to appreciate a broader role for NMD in the overall regulation of gene
expression. Given this, it has become increasingly important to understand the
underlying mechanisms of the NMD pathway and how an mRNA is able to trigger
degradation by NMD.
Aside from the presence of a PTC in an aberrant mRNA, there are several known
features that can target a wild-type mRNA for degradation by NMD: 1) a long 3’ UTR
(Amrani et al., 2004; Kebaara and Atkin, 2009; Muhlrad and Parker, 1999), 2) translation
of an upstream open reading frame (uORF; Amrani et al., 2006; Barbosa et al., 2013;
Nyiko et al., 2009), 3) a start codon in a suboptimal context which can lead to leaky
scanning and out of frame initiation of translation (Welch and Jacobson, 1999), 4) the
presence of programmed ribosome frameshift (PRF) sites (Plant et al., 2004), and 5) the
presence of pre-mRNA introns and regulated alternative splicing resulting in PTCs (He et
al., 1993; Lewis et al., 2003; McGlincy and Smith, 2008; Ni et al., 2007). All of these
NMD-targeting signals can result in either the ribosome terminating in a context that is
different from a normal termination event (i.e. proximity to the poly(A) tail; signals 1-2)
or in the introduction of a PTC (signals 2-4). Comparing wild-type mRNAs that have
NMD-targeting signals and are degraded by NMD with those that have NMD-targeting
signals and are not degraded by NMD may provide important insights as to the
underlying mechanisms of NMD.
Currently, there are four known mechanisms that can protect a wild-type mRNA
that has an NMD-targeting signal from degradation by NMD, but there are likely more to
50
be discovered. These NMD-protecting mechanisms include: 1) inhibition of translation,
2) translational readthrough (also known as stop codon suppression), 3) reinitiation of
translation, and 4) trans-acting factors (Carter et al., 1995; Dang et al., 2009; Keeling et
al., 2004; Neu-Yilik et al., 2011; Noensie and Dietz, 2001; Ruiz-Echevarria and Peltz,
2000; Wang et al., 2001). In support of the idea that there are more NMD-protecting
mechanisms yet to be discovered, we have identified an mRNA, SSY5, in Saccharomyces
cerevisiae that has multiple NMD-targeting signals but is protected from decay by the
NMD pathway through a novel mechanism.
The SSY5 mRNA codes for an essential component of the SPS amino acid sensor
in S. cerevisiae. The SPS sensor consists of three subunits: integral membrane protein
Ssy1, and peripheral membrane proteins Ssy5 and Ptr3. When extracellular amino acids
are present a signaling cascade is initiated by the binding of an amino acid to Ssy1, which
then transduces a signal via Ptr3 to Ssy5. Ssy5 is a serine protease that consists of two
domains: an inhibitory N-terminal pro domain and a catalytic C-terminal cat domain.
During the biogenesis of Ssy5 the two subunits are autolytically cleaved but remain
noncovalently associated until an activation signal is received (Andreasson et al., 2006;
Conrad et al., 2014; Ljungdahl, 2009). Once activated the Ssy5 pro domain is degraded
by the proteasome and the cat domain cleaves the N-terminal domain of transcription
factors Stp1 and Stp2, which allows them to enter the nucleus and activate the expression
of the amino acid permease (AAP) genes (Ljungdahl, 2009). AAP gene products then
facilitate the uptake of extracellular amino acids (Fig 1). Given the importance of Ssy5
51
and the intricacies of the SPS amino-acid-sensing pathway, it is likely that the regulation
of SSY5 mRNA levels is tightly controlled and has physiological significance.
Here we show the wild-type SSY5 mRNA, which has multiple NMD-targeting
signals, is not degraded by NMD. Based on previous studies, at least one of the targeting
signals—the long 3’ UTR—is able to target a reporter mRNA for degradation by NMD
(Obenoskey et al., 2014). We demonstrate that many of the current mechanisms for
protection of an mRNA from NMD do not apply to the SSY5 mRNA. This provides
evidence that the SSY5 mRNA is actually protected from degradation by NMD through
an unknown mechanism. Thus, the SSY5 mRNA provides a unique and intriguing case to
help elucidate the underlying mechanisms of gene regulation by the NMD pathway.
Materials and Methods
Yeast strains
The yeast strains used in this study are listed in Table 2-1. All yeast
transformations were done using Lithium Acetate-Mediated transformation as previously
described (Gietz and Woods, 2002). AAY590 was constructed by transforming AAY538
with the upf1Δ2 fragment from pAA70 using primers oAA48 and oAA79.
5’-tagged SSY5 mRNA constructs: AAY561 was constructed by transforming
BY4743 with the fragment amplified from the pFA6a-His3MX6-PGAL1-GFP plasmid
using primers F4 and R5 containing sequence to target the product to the SSY5 mRNA 5’
leader (Longtine et al., 1998). AAY568 was constructed by sporulation and random
spore analysis of AAY561. AAY625 was constructed by transforming AAY568 with the
52
upf1Δ2 fragment from pAA70 using primers oAA48 and oAA79. AAY630 was
constructed by transforming BY4743 with the fragment amplified from the pFA6aHis3MX6-PGAL1 plasmid using primers F4 and R2 containing sequence to target the
product to the SSY5 mRNA 5’ leader to make AAY559 (Longtine et al., 1998). AAY630
was the result of the sporulation and random spore analysis of AAY559. AAY632 was
constructed by transforming AAY630 with the upf1Δ2 fragment from pAA70 using
primers oAA48 and oAA79.
3’-tagged SSY5 mRNA constructs: AAY572 was constructed by transforming
BY4741 with the fragment amplified from pFA6a-GFP(S65T)-HIS3MX6 plasmid using
primers F2 and R1 containing sequence to target the product in-frame just before the
SSY5 mRNA stop codon (Longtine et al., 1998). AAY623 was constructed by
transforming AAY572 with the upf1Δ2 fragment from pAA70 using primers oAA48 and
oAA79. AAY581 was constructed by transforming BY4741 with the fragment amplified
from pFA6a-GFP(S65T)-HIS3MX6 plasmid using primers F2 and R1 containing
sequence to target the product in-frame just before the SSY5 mRNA stop codon that
would be used if reinitiation were to occur in the 3’UTR (Longtine et al., 1998).
AAY585 was constructed by transforming BY4741 with the fragment amplified from
pFA6a-GFP(S65T)-HIS3MX6 plasmid using primers F2 and R1 containing sequence to
target the product in-frame just before the SSY5 mRNA stop codon that would be used if
readthrough were to occur (Longtine et al., 1998). AAY572, AAY581 and AAY585
were sequenced to verify that the GFP sequence was inserted into the correct reading
frame.
53
SSY5 mRNA terminator constructs: AAY576 was constructed by transforming BY4741
with the fragment amplified from pFA6a-GFP(S65T)-HIS3MX6 plasmid using primers
F3 and R1 containing sequence to target the product to the SSY5 mRNA stop codon.
AAY601 was constructed by transforming AAY576 with the upf1Δ2 fragment from
pAA70 using primers oAA48 and oAA79.
Growth conditions
Unless otherwise noted yeast cells were grown using standard techniques with
mild agitation equivalent to 225rpm at 30˚C. When cells are stated as being grown in a
certain type of media, the same media was used to grow plate cultures from frozen stock
and all subsequent liquid cultures. In this study, YAPD media consists of: 1% yeast
extract, 2% Bacto-Peptone, 2% dextrose, and 100mg/L Adenine hemisulfate salt; YAPG
media consists of: 1% yeast extract, 2% Bacto-Peptone, 2% galactose, and 100mg/L
Adenine hemisulfate salt; and minimal media (SD+amino acids) consists of: 0.67% yeast
nitrogen base without amino acids, 2% dextrose, 20 mg/L L-Histidine, 30 mg/L LLeucine, 20 mg/L L-Methionine, 20 mg/L Uracil, and 30 mg/L L-Lysine.
Utilization of the GWIPS-viz Ribo-seq Genome Browser
Ribosome profiles were obtained from the QWIPS-viz Ribo-seq Genome
Browser: gwips.ucc.ie (Michel et al., 2014). Search: Group [Yeast], Genome [S.
cerevisiae], Assembly [Apr. 2011 (SacCer_Apr2011/SacCer3)], Position [chrX: 128,943128,987]. Select options: Reverse, then PS/PDF [Image configuration: text size-18,
54
image width-1346 pixels, label area width-14 characters, uncheck option to show light
blue vertical lines], download current browser in PDF.
RNA Extractions
Yeast strains were grown in 10mL cultures to an OD600 of 0.4-0.6. Cells were
harvested by centrifugation, washing in DEPC-ddH20, and flash-freezing in dry
ice/ethanol or liquid nitrogen. Cell pellets were stored at -70˚C until used for RNA
extractions. RNA extractions were performed as previously described (Kebaara et al.,
2012). RNA samples were diluted to 1µg/µl in DEPC-ddH20 and stored at -70˚C. RNA
quality check gels are performed for every RNA sample (1µl of 1µg/µl Total RNA is run
through a 0.8% agarose gel to check for degradation).
Quantitative Northern Analysis
10µg of Total RNA mixed with 3µl Formaldehyde loading dye (Ambion, cat. no.
8552) was separated through a 1.0% agarose gel containing 5.6% Formaldehyde and 1%
MOPS (10X MOPS: 0.2 M sodium morpholinopropanesulfonic acid (MOPS), pH 7.0,
0.05 M sodium acetate, 0.01 M EDTA; adjust pH to 7.0 with 10 M NaOH, do not
autoclave, store at room temperature in the dark.) RNA was transferred onto a
GeneScreen Plus membrane (PerkinElmer) using NorthernMax transfer buffer (Ambion,
cat. no. 8672) following the manufacturer’s protocol for downward transfer. The lane
with the RNA ladder was cut off of the gel before transfer and stained overnight in
55
0.5µg/ml ethidium bromide. Transfer was allowed to proceed for 2 hours with a 9mm
thick RNA gel. Membranes were rinsed in 2X SSC and dried for 15 minutes at 80˚C.
Membranes were hybridized with NorthernMax prehybridization/hybridization
buffer (Ambion # 8677).
32
P-labelled probes were synthesized using ~25ng of PCR
product corresponding to the gene of interest, the RadPrime DNA Labeling system
(Invitrogen #18428-011), and ∼50 μCi [α-32P]dCTP (3000 Ci/mmol, 10 mCi/ml) (Perkin
Elmer) following the manufacturer’s protocol. Probes are purified through a Sephadex
G-50 column equilibrated with TE pH 8.0. Membranes were hybridized overnight (12-24
hours) and then washed once at room temperature with 2X SSPE and once at 65˚C with
2X SSPE/2% SDS. Membranes were PhosphorImagedTM (GE Healthcare, Typhoon FLA
9500) and quantified using the ImageQuantTM software. All membranes were also
autoradiographed using a phosphorescent ruler to determine the size of the bands by
comparison to the RNA ladder. Membranes are stripped and stored at -20˚C for reprobing. Detailed protocol for Northern analysis can be found in Kebaara et al. (2012).
Westerns
Total protein was extracted as described in Atkin et al., 1995 from 40mls of midlog culture (OD600 of 0.4-0.6). Protein extracts were quantified in triplicate using BSA
standards (Pierce #23225). 40µg of total protein was separated through a 7.5% SDSPolyacrylamide gel and transferred to a .45µM Nitrocellulose membrane. Blocking was
done using 5% w/v Carnation® Instant Dry Milk overnight at 4˚C. Primary GFP
antibody used was a rabbit polyclonal ChIP-grade antibody (Abcam #ab290) and was
56
incubated at a concentration of 1:2000 diluted in 5% w/v Carnation® Instant Dry Milk
for 1hr at room temperature (per Abcam protocol). Secondary antibody was anti-rabbit
IgG, HRP-linked (Cell Signaling #7074) and was used at a concentration 1:6000 diluted
in 5% w/v Carnation® Instant Dry Milk for 1hr at room temperature (per Cell Signaling
protocol). Pierce ECL Western Blotting Substrate kit (Pierce #32109) was used for
detection. Signal was detected using film.
Polyribosome Analysis
Yeast polyribosome analysis was performed as previously described (Atkin et al.,
1995). Lysis buffer was composed of 20mM Tris-HCl pH 8.0, 140mM KCl, 1.5 mM
MgCl2, 1% Triton X, 0.1 mg/ml Cycloheximide, and 1.0 mg/ml Heparin; Cycloheximide
and Heparin were made fresh and added just before use. Gradient buffer (50% and 15%
sucrose) was composed of 20mM Tris-HCl pH 8.0, 140mM KCl, 5.0 mM MgCl2, 0.1
mg/ml Cycloheximide, 0.5 mg/ml Heparin, and 0.5mM DTT; Cycloheximide, Heparin,
and DTT were made fresh and added just before use. 15%-50% sucrose gradients were
made by hand, frozen at -70˚C and thawed at 4˚C overnight just before use. After
fractionation RNA was extracted from each fraction using acid phenol/chloroform
extraction. Total RNA from each fraction collected was loaded onto an RNA Northern
gel and transferred to a GeneScreen Plus membrane as described above. Membranes
were hybridized with a 32P-labelled probe as described above.
57
Results:
SSY5 mRNA has multiple NMD-targeting signals
The wild-type S. cerevisiae SSY5 mRNA was previously identified as an mRNA
with a long 3’ UTR that is not degraded by the NMD pathway (Kebaara and Atkin,
2009). Upon further inspection, we identified three additional potential NMD-targeting
signals present in the SSY5 mRNA: a translated uORF, a start codon in a poor context,
and five predicted ribosome frameshift sites (Fig. 2A-B; Belew et al., 2008; Guan et al.,
2006; Michel et al., 2014).
A map of ribosome footprints in the SSY5 uORF was compiled using the
GWIPS-viz Ribo-seq Genome Browser (Michel et al., 2014). The compiled ribosome
profile (summary profile from all studies) for the SSY5 mRNA uORF is displayed in
Figure 2B. The reading frame for the uORF is identified in the middle row (): the ATG
start codon begins at position 128, 986 (-34) and the UAA stop codon ends at position
128, 966. The reading frame for the SSY5 mRNA open reading frame (ORF) is identified
in the bottom row () with the ATG start codon beginning at position 128, 952 (+1).
The ribosome density peaks (in red) in the SSY5 mRNA 5’ leader region shows
ribosomes are present in a periodic pattern that represents translation (Ingolia et al.,
2009). When the ribosome terminates translation at the end of the uORF this termination
event can be recognized by the cellular machinery as a premature termination event,
which would target the mRNA for degradation by NMD (Guan et al., 2006; Hurt et al.,
2013).
58
Targeting to NMD by leaky scanning due to poor translation initiation at the start
codon is also a likely possibility for the SSY5 mRNA. Previously, H. Miyasaka
developed a method for quantifying the surrounding context of an AUG codon
(Miyasaka, 1999). This value, the AUGCAI (which stands for start Codon Adaptation
Index), is built on the concept of the CAI devised by Sharp and Li in 1987 and is used to
describe the optimal context of an AUG start codon (Miyasaka, 1999; Sharp and Li,
1987). The AUGCAI takes into account the nucleotide frequency at each position
surrounding the AUG from -6 to +6. Guan et al., (2006) revised the AUGCAI to test the
leaky scanning model for NMD. They created the AUGCAI(r) (start Codon Adaptation
Index revised), which is a score from 0 to 1.0 that takes into account both the frequency
at which each nucleotide appears at each position from -6 to +6 and how much each
nucleotide contributes to the overall start codon context, in order to determine the
efficiency of translation initiation at a particular AUG (Guan et al., 2006). Scores closer
to 0 indicate a start codon in a poor context that is more likely to be subject to leaky
scanning. Optimal start codon context, defined as (A/U)A(A/C)AA(A/C)AUGUC(U/C),
is assigned a value of 1.0; however, the frequency of mRNAs with an AUGCAI(r)=1.0 is
very low (data not shown; Hamilton et al., 1987). The SSY5 mRNA has an
AUGCAI(r)=0.375. In comparison, the AUGCAI(r) of the SPT10 mRNA, which is
targeted to NMD by leaking scanning, is 0.320 (Guan et al., 2006; Welch and Jacobson,
1999). Based on this observation, Guan et al. came to the conclusion that the best
candidates for leaky scanning are those with an AUGCAI(r) value ≤0.320. While the
AUGCAI(r) of the SSY5 mRNA is slightly above this cutoff, a value of 0.375 is still
59
relatively low and suggests that the start codon of the SSY5 mRNA could be subject to
leaky scanning.
Programmed ribosome frameshifting (PRF) can also cause an mRNA to become a
substrate for NMD. PRF sites are cis-elements within an mRNA that cause a ribosome to
shift reading frames either in the +1 direction or the -1 direction, although the -1 PRF is
much more frequent (Jacobs et al., 2007; Plant et al., 2004). A -1 PRF site is identified
by the presence of a heptameric “slippery site” sequence (X XXY YYZ, where X is any
three identical nucleotides, Y is either three A nucleotides or three U nucleotides, and Z
is either an A, U, or C) followed within eight nucleotides by a sequence that creates an
RNA pseudoknot (Dinman et al., 1991; Somogyi et al., 1993; Tu et al., 1992). A
previous study found that at least 2.54% of S. cerevisiae genes contain a consensus
sequence for a -1 PRF site. In contrast to viruses—where the PRF phenomenon was
originally discovered—the analysis in this study revealed that most of the -1 PRF signals
in yeast would result in the introduction of a PTC into the reading frame (Hammell et al.,
1999). We used the Programmed Ribosome Frameshift Database (PRFdb), a database
that contains computationally predicted -1 PRF sites in eukaryotic genomes, to analyze
the SSY5 mRNA for the presence of -1 PRF sites (Belew et al., 2008). The PRFdb
predicts five potential -1 PRF sites in the SSY5 mRNA at +18, +132, +324, +333, and
+1728. Using the GWIPS-viz Genome Browser, which shows all three reading frames of
an mRNA, we noted that any shift in the reading frame of the SSY5 mRNA results in the
introduction of multiple stop codons throughout the ORF (Michel et al., 2014). However,
the predicted -1 PRF sites in the SSY5 mRNA have not yet been experimentally validated.
60
Finally, long 3’ UTRs have been identified in both S.cerevisiae and humans as
NMD-targeting signals (Amrani et al., 2004; Eberle et al., 2008; Hogg and Goff, 2010;
Kebaara and Atkin, 2009; Muhlrad and Parker, 1999; Rebbapragada and LykkeAndersen, 2009; Yepiskoposyan et al., 2011). One explanation for how PTCs trigger
NMD was proposed in the faux 3’ UTR model which suggests that early translation
termination (e.g. at a PTC) is distinct from normal termination because the terminating
ribosome is not in the proper context (e.g. proximity) to interact with 3’ UTR-bound
poly(A) binding protein, Pab1. The failed interaction between the terminating ribosome
and Pab1 leaves the ribosome open for binding of the NMD factors (Upf1, Upf2, and
Upf3), which triggers NMD. A long 3’ UTR would place the stop codon, even though it
is not a PTC, in the same context (i.e. too far from the poly(A) tail for the terminating
ribosome to bind Pab1) as if it were a PTC. In S. cerevisiae, most 3’ UTRs are short
ranging in length from 50-200 nt with a median length of 121nt (Graber et al., 1999).
The SSY5 mRNA has a 3’ UTR of ~475 nt determined by 3’ RACE (Obenoskey et al.,
2014). The use of alternative Poly(A) sites is predicted to produce SSY5 mRNA 3’UTRs
that range from 420-500nt (Kebaara and Atkin, 2009).
The long 3’ UTR of the SSY5 mRNA has been validated as an NMD-targeting
signal by replacing the native 3’ UTR of the CYC1 mRNA, which is normally NMDinsensitive, with the long 3’ UTR of the SSY5 mRNA. The CYC1-SSY5 3’UTR construct
is a substrate for NMD indicating that the long 3’ UTR of the SSY5 mRNA is sufficient to
target an mRNA for degradation by NMD (Obenoskey et al., 2014). However, the
61
remaining three potential NMD-targeting signals (Fig. 2A 1-3) have yet to be validated as
positive NMD-targeting signals for the SSY5 mRNA.
Codon optimality of an mRNA was recently identified as a significant
determinant of mRNA stability (Presnyak et al., 2015). A higher composition of codons
designated as “optimal” results in stable mRNAs whereas a lower composition results in
unstable mRNAs and higher turn-over. The optimal codon content of an mRNA also
influences the elongation rate of the translating ribosome where a lower content of
optimal codons results in a slower elongation rate (Presnyak et al., 2015). The optimal
codon content of SSY5 mRNA is 285 optimal codons out of 698 codons total (subtracting
out the AUG start codon and UGA stop codon) yielding a 40.83% optimal codon content.
The original study showed that mRNAs with less than 40% optimal codon content are
significantly more unstable (median half-life of 5.3 min) than mRNAs with 70% optimal
codon content (median half-life of 20.1 min; Presnyak et al., 2015). Thus, SSY5 mRNA
percent optimal codon content falls much unstable mRNA category than to the stable
mRNAs. However, it should be noted that codon optimality is subject to exceptions and
the contribution of codon optimality specifically to the stability of the SSY5 mRNA
remains to be experimentally determined.
The SSY5 mRNA has four of the known NMD-targeting features: 1) a translated
uORF, 2) a start codon in a poor context based on the AUGCAI(r) value of 0.375, 3) five
predicted -1 PRF sites that would all introduce multiple downstream stop codons, and 4)
a long 3’ UTR that would result in the ribosome terminating a long distance from the
poly(A) tail. Additionally, the percent of optimal codons that compose the SSY5 mRNA
62
is low, and that alone should destabilize the mRNA. All of this leads to the rational
assumption that the SSY5 mRNA should be a substrate for degradation by NMD.
However, below we show this is not the case.
SSY5 mRNA is not degraded by NMD
An initial screen of mRNA half-lives (T1/2) in wild-type and upf1Δ strains
identified the SSY5 mRNA as being unaffected by the NMD pathway (Kebaara and
Atkin, 2009). In order to validate these results we grew S.cerevisiae cells in rich growth
medium (YAPD) and confirmed by quantitative Northern analysis that steady-state SSY5
mRNA levels show no significant fold change (FC) between wild-type and upf1Δ strains
(fold change ratio (FCR) of upf1Δ/ wild-type =1.2±0.1) in the BY4741 genetic
background (Fig. 3A). Further, the T1/2 of the SSY5 mRNA was not significantly
different between wild-type (T1/2=33.3min) and upf1Δ strains (T1/2=38.3min; Fig. 3B).
ENT4 is an mRNA with a long 3’ UTR (~600 nt) that was previously identified in S.
cerevisiae as a substrate for NMD (Kebaara and Atkin, 2009). Here we use the ENT4
mRNA as well as the CYH2 pre-mRNA as positive controls for NMD, and SCR1 serves
as the loading control.
It was previously shown that different genetic backgrounds in yeast can influence
the relative accumulation of nonsense mRNAs in wild-type versus upf1Δ strains to
varying degrees (Kebaara et al., 2003). Thus, steady-state SSY5 mRNA levels were also
determined in wild-type and upf1Δ strains in the W303 genetic background. Consistent
with results in the BY4741 background, there was no significant fold change in steady-
63
state SSY5 mRNA levels between wild-type and upf1Δ strains (FCR of upf1Δ/wild-type
=1.1±0.1) in the W303 background (Fig. 3A). Half-lives were not determined for the
W303 background in the present study as these were shown in a previous study (Kebaara
and Atkin, 2009). The BY4741 and W303 genetic backgrounds are two of the most
commonly used genetic backgrounds in NMD work with S. cerevisiae. BY4741 strains
are derived from S288C and differences between the two are very minor (Brachmann et
al., 1998). W303 is a lab strain developed as a model organism and differs from S288C
at >8,000 nucleotide positions, but still shares >85% of its genome with S288C (Ralser et
al., 2012). Based on these results all subsequent experiments were done using strains
with the BY4741 background.
The SSY5 mRNA does not appear to be a substrate for NMD when cells are
grown in rich media (Fig. 3A-B). However, since the SSY5 mRNA codes for the catalytic
subunit of an amino-acid-sensing complex (Fig. 1), we hypothesized that the stability of
the SSY5 mRNA might vary in different growth conditions. To test this hypothesis we
grew the cells in minimal media which contained only the amino acids necessary to
support the auxotrophies of the strains (Materials and Methods). The steady-state
accumulation of the SSY5 mRNA was not significantly different between wild-type and
upf1Δ strains (FCR=1.1±0.2; Fig. 3C). Additionally, the T1/2 of the SSY5 mRNA was not
significantly different between wild-type (T1/2=12.4min) and upf1Δ strains (T1/2=13.3min;
Fig. 3D). Although the T1/2 of the SSY5 mRNA is not significantly different between
wild-type and upf1Δ strains in both types of media (rich vs. minimal) we did note that the
overall T1/2 of the SSY5 mRNA in both strains (wild-type and upf1Δ) is much longer in
64
rich media than in minimal media (Fig. 3B vs. Fig. 3C). This is likely due to the SPSsensor being in an active state during growth in rich media leading to higher turnover of
the Ssy5 component. An increase in demand for Ssy5 could lead to increased mRNA
T1/2.
Typically, NMD substrates are rapidly degraded in wild-type cells and
accumulate to significantly higher levels in upf1Δ strains. Additionally, the T1/2 of NMD
substrates is significantly longer in upf1Δ strains compared to wild-type strains (Cao and
Parker, 2003; He and Jacobson, 2001; Muhlrad and Parker, 1999). Based on our results
for steady-state mRNA accumulation and T1/2 analysis of the SSY5 mRNA in both rich
and minimal media we conclude that the SSY5 mRNA is not degraded by the NMD
pathway and this stability is not attributed to differences in environmental amino acid
content. However, it should be noted that the minimal media used still contains five
amino acids, so true amino acid starvation was not obtained due to auxotrophic
requirements of the strains (Materials and Methods).
Replacing the long 3’UTR of SSY5 mRNA with a short 3’UTR does not influence
SSY5 mRNA susceptibility to NMD
Given that the SSY5 mRNA has multiple NMD-targeting signals (Fig. 2), and that
at least one of these targeting signals (the long 3’ UTR) is sufficient to target an mRNA
for degradation by NMD (Obenoskey et al., 2014), yet we do not find an increase in
steady-state accumulation of SSY5 mRNA or a difference in T1/2 between wild-type and
upf1Δ strains (Fig. 3A-D) we hypothesize that the SSY5 mRNA is somehow protected
65
from degradation by the NMD pathway. As a start to identifying the feature(s) and/or
region(s) of the SSY5 mRNA responsible for protection from NMD we began with the 3’
UTR. Since it has already been determined that the long 3’ UTR of the SSY5 mRNA,
when placed on a different ORF, is sufficient to target an mRNA to NMD, we began with
the reciprocal experiment of replacing the long 3’ UTR of the SSY5 mRNA with a short
3’ UTR (Obenoskey et al., 2014). To do this we selected the 180nt 3’ UTR of the ADH1
mRNA (Fig. 4A). The ADH1 mRNA 3’ UTR was selected for two reasons: 1) it has a
length that falls within the range of typical S. cerevisiae 3’ UTRs (50-200nt), and 2) the
tools for replacing the 3’ UTR of an mRNA with the ADH1 3’ UTR are readily available
and have been validated (Graber et al., 1999; Longtine et al., 1998).
Using quantitative Northern analysis, we found no significant difference in
steady-state mRNA accumulation of the SSY5+ADH1 3’ UTR mRNA between wild-type
and upf1Δ strains (FCR=1.1±0.2; Fig. 4B, lanes 2 and 3). Likewise, there was no
significant increase in T1/2 of the SSY5+ADH1 3’ UTR mRNA in the upf1Δ strain
(T1/2=16.3min) compared to the wild-type strain (T1/2=12.3min; Fig. 4C). However, we
did observe a higher accumulation of SSY5+ADH1 3’ UTR mRNA in both wild-type and
upf1Δ strains compared to the normal SSY5 mRNA (FCR=1.6±0.2 and FCR=1.7±0.0,
respectively; Fig. 4B, compare lanes 2 and 3 with lane 1).
Additionally, because SSY5 mRNA codes for an essential component of the SPS
amino-acid-sensing complex we wanted to see if altering the 3’ UTR of the SSY5 mRNA
had any influence on the function of Ssy5. We considered this because sequence
elements lead to speculation that readthrough of the annotated SSY5 mRNA stop codon is
66
a possibility (discussed below, Fig. 7). If readthrough is occurring, then it is possible that
replacing the native 3’ UTR of the SSY5 mRNA could influence Ssy5 structure and
function. To determine if Ssy5 enzymatic function is compromised in this strain we
chose to look at mRNA accumulation of a downstream target of Ssy5. Ssy5 is
responsible for the proteolytic processing of the transcription factors Stp1 and Stp2,
which, after being cleaved by Ssy5, enter the nucleus and upregulate the expression of
amino acid permease (AAP) genes. The AGP1 mRNA codes for an amino acid
transporter with broad substrate range and is one of the AAP genes upregulated in
response to Stp1/2 cleavage by Ssy5 (Fig.1; Ljungdahl, 2009). There is no significant
difference in the accumulation of the AGP1 mRNA between the strain with the native
SSY5 3’ UTR (Fig. 4B, lane 1) and the strains with the SSY5+ADH1 3’ UTR construct
(Fig. 4B, lanes 2 & 3). Thus, using AGP1 mRNA accumulation as an indicator of Ssy5
activity, there is no evidence of altered Ssy5 function as a result of replacing the SSY5
mRNA 3’ UTR with the shorter ADH1 mRNA 3’ UTR.
Taken together, these results show that replacing the long 3’ UTR of the SSY5
mRNA with a shorter 3’ UTR has no impact on SSY5 mRNA stability. Hence, we
conclude that the 3’ UTR of the SSY5 mRNA is not responsible for the protection of SSY5
mRNA from NMD. This is also significant because a previous study identified a
stabilizer element (STE) that is able to protect mRNAs from NMD (Ruiz-Echevarria and
Peltz, 2000). This study showed that the STE must be downstream of a stop codon to
confer protection from NMD. Here we removed all the SSY5 3’ UTR sequence
downstream of the stop codon, so even if there was an STE in the 3’ UTR that is not the
67
mechanism of protection of the SSY5 mRNA from NMD. However, the long 3’ UTR is
not the only NMD-targeting signal present in the SSY5 mRNA. Investigation of the
additional targeting signals may provide insight as to the mechanism of protection of the
SSY5 mRNA from NMD.
Perturbation of the SSY5 mRNA 5’ UTR does not affect SSY5 mRNA stability
Another potential NMD-targeting feature present in the SSY5 mRNA is the
presence of a translated uORF (Fig. 2B). Since replacing the long 3’ UTR of the SSY5
mRNA with a shorter 3’ UTR did not provide evidence of a stabilizing element in the 3’
UTR we decided to refocus our attention to the uORF. For this, we hypothesize that if
disruption of the 5’ UTR influences SSY5 mRNA stability then there are two likely
possibilities: 1) there is another cis-acting element present within the 5’ UTR responsible
for SSY5 mRNA stability, or 2) there are trans-acting factors that bind to specific
sequences within the 5’ UTR to confer stability. It is also possible that SSY5 mRNA
stability could be the result of a combination of any of these three ideas.
To determine if the SSY5 mRNA stabilizing element is located in the 5’ UTR we
utilized a construct that had already been generated in our lab. This construct uses a
previously validated method for replacing the 5’ UTR of an mRNA with the S. cerevisiae
galactose (GAL1-10) promoter sequence (pGAL; Longtine et al., 1998). Using this
method, 50nt of the SSY5 5’ UTR are replaced with the pGAL sequence, thus abolishing
the SSY5 uORF and the majority of the SSY5 5’ UTR sequence.
68
Using quantitative Northern analysis we measured the steady-state accumulation
of the pGAL-SSY5 mRNA in wild-type and upf1Δ strains. No significant difference in
accumulation between the two strains (FCR=1.1±0.1) was observed. We also measured
the pGAL-SSY5 mRNA T1/2 in wild-type and upf1Δ strains. Although there is a slight
increase in pGAL-SSY5 mRNA T1/2 in the upf1Δ strain (T1/2=57.6min) compared to wildtype strain (T1/2=41.6min) this difference is too small to be considered significant (Fig
5B). If the pGAL-SSY5 mRNA does become a substrate for NMD then we would expect
to see ≥2-fold increase in T1/2 in the upf1Δ strain. These results indicate that abolishing
the native SSY5 5’ UTR sequence does not influence SSY5 mRNA stability, thus the 5’
UTR does not contain the stabilizing element.
An important consequence of disruption of the SSY5 5’ UTR using the above
method is that SSY5 mRNA is now under the control of an inducible promoter. Using
this we can also determine the effects of overexpression of the SSY5 mRNA. When SSY5
mRNA is placed under the control of the galactose promoter and the cells are grown in
rich media containing 20% galactose (YAPG) we can achieve ~65-fold increase in SSY5
mRNA expression (Fig. 5C). Significantly, this substantial increase in SSY5 mRNA
expression has no influence on SSY5 mRNA stability (Fig. 5A and B) and also has no
consequence on Ssy5 downstream function (Figure 5C, AGP1 mRNA levels). This
indicates that significant overexpression of the SSY5 mRNA in the cell does not saturate
the stabilizing mechanism (e.g. trans-acting factors). It also indicates that there is a ratelimiting step before Ssy5 protease activity in the SPS sensor signaling cascade. Also
69
noted, the cells grow at average rates in liquid media and on plates, so overexpression of
the SSY5 mRNA is not toxic to the cells (data not shown).
SSY5 mRNA is not protected from NMD by inhibition of translation
NMD is a translation-dependent mechanism since targeting of an mRNA to NMD
typically involves the interaction of the core NMD factors with the terminating ribosome
(Bertram et al., 2001; Czaplinski et al., 1998; Kisselev et al., 2003; Rospert et al., 2005).
In line with this, several studies have shown that the inhibition of translation, by blocking
translation initiation or elongation, is able to suppress NMD (Carter et al., 1995; Dang et
al., 2009; Noensie and Dietz, 2001). We hypothesize that inhibition of translation is not
the stabilizing mechanism of the SSY5 mRNA because ssy5Δ mutants are not viable and
the Ssy5 protein is an essential component of the SPS sensor, which is required for proper
cell homeostasis (Conrad et al., 2014; Ljungdahl, 2009). Nevertheless, it is important to
confirm that the SSY5 mRNA is actively translated in order to rule out this possibility.
Polyribosome analysis was performed using cell extracts from wild-type and
upf1Δ strains. Density gradient centrifugation of total cell extracts through 15%-50%
sucrose gradients was performed followed by gradient fractionation. Total RNA was
extracted from each fraction and quantitative Northern analysis was used to determine the
mRNA distribution among fractions (Materials and Methods). SSY5 mRNA is found
primarily in the polyribosome fractions (9-15) of both wild-type and upf1Δ strains
indicating that the mRNA does co-migrate with polyribosomes in both strains, which
provides strong evidence in support of SSY5 mRNA association with polyribosomes (Fig.
70
6A). The polyribosome analysis of the CYH2 pre-mRNA and mRNA in wild-type and
upf1Δ strains is provided for comparison to show how the distribution of an NMDsubstrate (CYH2 pre-mRNA) differs from a non-NMD substrate (CYH2 mRNA) between
wild-type and upf1Δ strains (Fig. 6A). In comparison, the SSY5 mRNA distribution
appears to mirror that of the CYH2 mRNA, which is translated and not an NMDsubstrate. However, we cannot ignore the presence of the slower migrating SSY5 band
that is specific to the polyribosome fractions in the upf1Δ strain. This band could be a
longer or modified form of the SSY5 mRNA that is actually a substrate for NMD.
To confirm that SSY5 mRNA co-migration with polyribosomes is in fact
indicative of active translation we constructed a strain in which GFP is fused to the 5’end
of the SSY5 mRNA. Because endogenous levels of SSY5 mRNA and Ssy5 protein are
extremely low we again employed the pGAL promoter when constructing these strains.
GFP fused to the SSY5 mRNA under the control of the native promotor was not
detectable by fluorescence microscopy or flow cytometry (data not shown). By placing
the GFP-SSY5 mRNA under the control of the GAL promoter (pGAL-GFP-SSY5) we
were able to visualize GFP-Ssy5 using confocal fluorescence microscopy (Fig 6B). GFP
can be seen in the strains with the pGAL-GFP-SSY5 mRNA but not in the strains without
the GFP-tagged Ssy5 (Fig. 6B, compare top and bottom rows). Importantly, yeast cells
do exhibit green autofluorescence; however we were able to filter out this
autofluorescence (Fig. 6B, bottom row) so the green fluorescence seen in the pGAL-GFPSSY5 mRNA panels is specific to GFP-Ssy5. Hoechst was used as the nuclear stain.
These results confirm SSY5 mRNA is actively translated.
71
Furthermore, we confirmed that addition of the pGAL-GFP sequence does not
alter stability of the SSY5 mRNA as the pGAL-GFP-SSY5 mRNA accumulates to similar
levels in both wild-type and upf1Δ strains (FCR=1.1±0.1; Fig. 6C). This result is
consistent with the results observed in Figure 5 for the pGAL-SSY5 mRNA without the
GFP sequence.
The SSY5 mRNA 3’ UTR is a good candidate for both stop codon readthrough and
reinitiation of translation
Upon closer analysis of the SSY5 mRNA 3’ UTR we find two unique features that
are of interest concerning potential mechanisms of escape or protection of the mRNA
from NMD. First, the SSY5 mRNA stop codon is a good candidate for stop codon
readthrough. Second, the SSY5 mRNA 3’ UTR has the potential for translation
reinitiation. Either of these mechanisms independently would provide an explanation for
the stability of the SSY5 mRNA (Bonetti et al., 1995; Keeling et al., 2004; Neu-Yilik et
al., 2011; Wang et al., 2001).
Previously, it was shown that leaky termination at PTCs allows mRNAs to escape
degradation by NMD (Keeling et al., 2004). Following this idea, it would make sense
that leaky termination at the SSY5 stop codon could protect the SSY5 mRNA from NMD
as it would reduce the length of the SSY5 mRNA 3’ UTR bringing the terminating
ribosome in closer proximity to Pab1. There are three features of the SSY5 mRNA stop
codon and 3’ UTR that lead us to hypothesize there is a high probability of stop codon
readthrough. First, the SSY5 mRNA terminates with a UGA stop codon. In yeast, the
72
UGA stop codon is recognized less efficiently by the terminating ribosome than the UAA
or UAG stop codons (Keeling et al., 2004). Second, the SSY5 mRNA stop codon is in a
suboptimal context. Optimal stop codon context occurs when the stop codon is followed
immediately by a purine residue (A/G) whereas the SSY5 mRNA stop codon is followed
by a T (Fig. 7; Bonetti et al., 1995). Additionally, it has been shown that very efficient
termination occurs when the codon immediately upstream of the stop codon is a UCC
(serine) codon and immediately downstream is a GCA (alanine) codon, and the SSY5
mRNA contains neither of these (Bonetti et al., 1995; Namy et al., 2001). Third, the
SSY5 mRNA UGA stop codon is not immediately followed by another in-frame stop
codon or codons—a common feature of efficient termination context (Williams et al.,
2004). If the ribosome failed to terminate at the initial UGA stop codon translation
would proceed a significant distance (102 codons) before encountering another in-frame
stop codon. Moreover, the next downstream in-frame stop codon is a more efficiently
recognized stop codon (UAA) and is followed by a purine residue, A (Fig. 7; Keeling et
al., 2004).
Previous work on the human β-globin mRNA presented a puzzling group of
mutations in the first exon of the mRNA. Typically, nonsense mutations near the 5’ end
of the mRNA elicit degradation by NMD (Chamieh et al., 2008; Le Hir et al., 2001;
Lykke-Andersen et al., 2001; Shibuya et al., 2004; Singh et al., 2007). Interestingly,
there are a group of nonsense mutations in the first exon of the human β-globin mRNA
that do not trigger degradation by NMD (Inacio et al., 2004; Romao et al., 2000). Further
studies showed there is a sharp boundary in the position at which nonsense codons in this
73
mRNA trigger NMD or do not (Neu-Yilik et al., 2011). It was shown that the reason for
this sharp boundary and inability of the first exon nonsense mutations to trigger NMD
was due to reinitiation of translation at a downstream AUG (Neu-Yilik et al., 2011). The
SSY5 mRNA 3’ UTR has the likely potential for translation reinitiation as there is a start
codon not far downstream from the original stop codon but in a different reading frame.
This start codon is followed by an 82-codon ORF that terminates with a UAA termination
codon in a good context (i.e. followed by a purine residue; Fig. 7). The idea of
reinitiation in the mRNA 3’ UTR is intriguing as it introduces the possibility of a
regulatory downstream open reading frame (dsORF) that may serve similar functions as
the uORF.
The SSY5 mRNA is not protected from NMD by stop codon readthrough
We have identified reasons to speculate there is a good chance for stop codon
readthrough of the SSY5 mRNA UGA codon (discussed above). Thus, we hypothesize
that stop codon readthrough may be protecting the SSY5 mRNA from NMD. To test this
idea we designed a reporter system that is similar to the one used previously (Bonetti et
al., 1995). Briefly, we placed a GFP tag in-frame in the SSY5 mRNA 3’ UTR
immediately preceding the downstream UAA stop codon that would likely be used if
readthrough of the original UGA stop codon was to occur (Reporter #2; Fig. 8A, orange
arrow). As a positive control we also made a reporter construct where the GFP tag was
inserted in-frame immediately preceding the original SSY5 mRNA UGA stop codon
(Reporter #1; Fig. 8A, green arrow).
74
Total protein extracts were examined for the presence of GFP in strains
containing either reporter using Western analysis. Ssy5 has a molecular weight of 77.5
kDa while GFP has a molecular weight of 26.9 kDa, so the control reporter should appear
at 104.4 kDa. Reporter #2 contains the additional sequence of the SSY5 3’ UTR that
would be translated if readthrough were to occur which adds another 0.3 kDa to the total
protein size. We were able to detect a band in the positive control at ~104.4 kDa that was
not present in the negative control—untagged Ssy5 (Fig. 8B, compare lane one with lane
two top asterisk). However, we were not able to detect a band at ~104.7 kDa in the
readthrough reporter (Fig. 8B, compare lane two with lane three top asterisks).
Additionally, we looked for GFP signal in strains containing each construct using
fluorescence microscopy and flow cytometry. However, in these constructs each reporter
is under the control of the native SSY5 promoter so protein levels are extremely low and
we were unable to detect GFP signal even in the positive control using either method
(data not shown).
Using total protein extracts from the strains and a ChIP-quality GFP antibody we
still observed the presence of numerous non-specific cross-reacting bands that can be
seen in Figure 8 B. Conversely, there is one band that is specific to the protein extract
from the positive control (Fig. 8B, lane 2 bottom asterisk). In looking for bands at 104.4
kDa and 104.7 kDa we only accounted for the full-length uncleaved Ssy5-GFP.
However, during the biogenesis of Ssy5 the protein is autolytically cleaved into the 35.5
kDa catalytic C-terminal cat-domain and the 42.2 kDa inhibitory N-terminal pro-domain
(Andreasson et al., 2006). For the GFP reporter constructs used here we inserted the GFP
75
sequence at the 3’ end of the mRNA, which places the GFP tag at the C-terminal end of
the cat-domain. So, the band we observe at ~62.2 kDa is specific to Ssy5 and accounts
for the GFP-tagged cat-domain of Ssy5 that has been autolytically cleaved from the prodomain (Fig. 8B, lane 2 bottom asterisk and schematic on the right).
We also analyzed the steady-state mRNA levels in each construct to confirm that
addition of the 3’ GFP sequence in either location does not influence SSY5 mRNA
accumulation. As expected, the stead-state mRNA accumulation is similar between wild
type SSY5 mRNA and both SSY5-GFP mRNA constructs (Fig. 8C). Also, addition of the
3’ GFP tag does not appear to influence Ssy5 activity as downstream AGP1 mRNA levels
remain unchanged in the Ssy5-GFP constructs compared to the untagged Ssy5 (Fig. 8C).
Here we show that are able to detect Ssy5-GFP by Western analysis through the
presence of two specific bands: one at 104.4 kDa and one at 62.2 kDa. However, we are
unable to detect the presence of specific Ssy5-GFP bands using the reporter designed to
test for readthrough (Fig 8B). This leaves us with two possibilities: 1) readthrough of the
SSY5 mRNA UGA is not occurring or, 2.) SSY5 mRNA UGA readthrough is not
occurring at a high enough frequency for us to be able to detect a readthrough product.
Using the positive control reporter we determined that we cannot detect a specific Ssy5GFP band when ≤10µg of total protein extract is loaded. Based on these results, we can
conclude the SSY5 mRNA is not likely protected from NMD by stop codon readthrough.
76
The SSY5 mRNA is not protected from NMD by translation reinitiation
In addition to stop codon readthrough, we have also identified features of the
SSY5 mRNA 3’ UTR that are favorable for translation reinitiation (discussed above).
Therefore, we hypothesize that translation reinitiation in the SSY5 mRNA 3’ UTR may be
protecting the SSY5 mRNA from NMD. To test this idea we again used our GFP reporter
system designed above for the read through analysis. We used the same reporter for the
positive control (reporter #1 with GFP inserted in frame immediately preceding the SSY5
mRNA UGA stop codon; Fig. 9A, green arrow). The reporter designed to test for
translation reinitiation places GFP in-frame with the downstream open reading frame
immediately preceding the UAG stop codon that would be used for termination of this 3’
UTR ORF (Reporter #3; Fig. 9A, blue arrow).
Total protein extracts were examined for the presence of GFP in strains
containing either reporter using Western analysis as results using fluorescence
microscopy and flow cytometry proved unsuccessful because of the low level of protein
expression under the control of the native SSY5 mRNA promoter (data not shown). We
expected the same bands as before for the positive control at 104.4 kDa and 62.2 kDa.
The translation reinitiation product, however, is much smaller and should appear at ~
36.6 kDa. Since we did not separate the protein extracts using a gradient gel and the
reinitiation product is so small, we were only able to distinguish the protein product for
the processed GFP-tagged cat-domain in the positive control at 62.2 kDa because the
104.4 kDa was obscured by the group of cross-reacting bands (Fig. 9B). Nevertheless,
based on our results in Figure 8B this band still allows us to confirm specificity for an
77
Ssy5-GFP product. However, we were unable to detect a reinitiation product at 36.6 kDa
(Fig. 9B). Again, this means that either reinitiation in the SSY5 3’ UTR is not occurring
or that the reinitiation is occurring at a frequency too low to be detected by our methods.
Additionally, it is possible that reinitiation may be occurring in a different reading frame
with a different AUG, as there are five additional out-of-frame AUG codons. We tested
the most likely position for reinitiation to occur based on start codon context, size of the
ORF, and context of the stop codon.
We also analyzed steady-state SSY5 mRNA levels in all of the constructs to
confirm that addition of the 3’ UTR GFP sequence to test for translation reinitiation did
not alter mRNA accumulation (Fig 9C). The small decrease in accumulation in the SSY5GFP reinitiation mRNA construct is not considered significant. The more diffuse
banding pattern for the SSY5 mRNA in all of the 3’ GFP-tagged constructs in noted, and
is likely the result of a more diverse population of mRNAs, but with small variation (Fig,
8C and Fig. 9C). Furthermore, addition of the 3’ UTR GFP sequence in the reinitiation
reading frame does not appear to influence Ssy5 activity as downstream AGP1 mRNA
levels remain unchanged in the Ssy5-GFP constructs compared to the untagged Ssy5
(Fig. 9C).
Thus, we conclude that the SSY5 mRNA is most likely not being protected from
NMD due to translation reinitiation in the SSY5 3’ UTR at the most favorable potential
downstream open reading frame.
78
SSY5 mRNA is translated equally efficient in both wild-type and upf1Δ cells
The current literature provides evidence that if translation initiation is inhibited
then an mRNA is stable and not a substrate for degradation by NMD (Dang et al., 2009;
Low et al., 2005; Zuk and Jacobson, 1998). Additionally, translation elongation
inhibitors such as cycloheximide and puromycin also block degradation of substrates by
NMD (Carter et al., 1995; Herrick et al., 1990; Noensie and Dietz, 2001; Zhang et al.,
1997). Finally, if translation proceeds past the ORF stop codon (i.e. stop codon
readthrough/nonsense suppression) then NMD is also inhibited and the mRNA is stable
(Keeling et al., 2004). Further, mutations in any of the three NMD trans-acting factors
(Upf1, Upf2, or Upf3) effectively reduce the efficiency of translation termination
(Maderazo et al., 2000; Wang et al., 2001). Importantly, it was previously shown that
recognition of a nonsense codon in an mRNA leads to an overall decrease in the
translational efficiency of the mRNA as a reporter carrying a nonsense mutation yielded
significantly higher levels of protein in upf1Δ strains compared to wild-type (Muhlrad
and Parker, 1999).
Together this information led us to the hypothesis that if the SSY5 mRNA is being
recognized as an NMD substrate, and the mRNA is protected at a point after recognition,
then inactivation of the NMD pathway (upf1Δ strains) should show significantly higher
Ssy5 levels compared to wild-type strains. To test this hypothesis we utilized our SSY5GFP reporter strains that were constructed above (Fig. 8 and Fig 9). The reporter
construct containing the GFP tag in-frame just before the SSY5 mRNA stop codon
(Reporter #1) was transformed into NMD+ (wild-type strains used previously) and nmd-
79
(upf1Δ) strains. Total protein extract was analyzed for Ssy5-GFP, which should produce
bands at 104.4 kDa and 62.2 kDa (Fig. 10, lanes 2 and 3 asterisks). Nonspecific crossreacting bands in all samples are shown to verify loading. We were able to detect the
specific Ssy5-GFP product in the NMD+ and nmd- strains, however, there does not
appear to be a significant difference in the amount of protein present between the two
strains (Fig. 10, lanes 2 and 3 asterisks). These results indicate that the NMD pathway is
not likely playing a role in the translation of the SSY5 mRNA. Thus, the SSY5 mRNA is
probably not being recognized as an NMD substrate, which is consistent with the rest of
our results.
SSY5 mRNA is not protected from NMD by RNA binding protein Pub1
Finally, we wanted to determine if SSY5 mRNA is protected from NMD due to
the interaction of trans-acting factors. The search for potential candidates is a
cumbersome task as the pool RNA binding proteins (RBPs) is quite large and there is no
collective database that identifies specific binding motifs for the different RBPs.
Additionally, it may be an indirect interaction that could be responsible for SSY5 mRNA
protection from NMD in which case identifying RBP binding motifs present in the SSY5
mRNA would be uninformative. However, a recent study showed that poly(U) binding
protein (Pub1) is able to protect two wild-type mRNAs, GCN4 and YAP1, from
degradation by NMD (Ruiz-Echevarria and Peltz, 2000). These two mRNAs are targeted
for degradation by NMD through the presence of translated uORFs. It was shown that
both GCN4 and YAP1 mRNAs contain stabilizer elements (STEs) in their 5’ UTR which
80
binds Pub1. The binding of Pub1 protects the mRNA from degradation by NMD (RuizEchevarria and Peltz, 2000).
Using this information we hypothesized that Pub1 could play a role in SSY5
protection from NMD. To determine if Pub1 is protecting SSY5 mRNA from degradation
by NMD, we analyzed steady-state SSY5 mRNA accumulation in wild-type, upf1Δ,
pub1Δ, and upf1Δpub1Δ strains when cells are grown in rich media (YAPD).
Quantitative Northern analysis reveals no significant differences in SSY5 mRNA
accumulation in pub1Δ (FCR of pub1Δ/wild-type =1.0±0.2) or upf1Δ pub1Δ (FCR of
upf1Δpub1Δ/wild-type =1.2±0.2) cells compared to wild-type cells (Fig. 11A). We also
tested the possibility that Pub1 could be protecting SSY5 mRNA only under nutrientlimiting conditions by analyzing steady-state SSY5 mRNA accumulation in wild-type,
upf1Δ, and pub1Δ strains when cells are grown in minimal media (amino-acid limiting).
We observed the same trend in this condition with no significant difference in SSY5
mRNA accumulation in pub1Δ (FCR of pub1Δ/wild-type =1.0±0.2) cells compared to the
wild-type cells (Fig. 11B).
Further, we confirmed that there is no change in SSY5 mRNA stability in wildtype vs pub1Δ cells by determining the T1/2 of SSY5 mRNA in each of the strains. The
SSY5 mRNA had a T1/2 of 10.2±1.2 min in wild-type strains compared to a T1/2 of
14.6±1.7 min in pub1Δ (Fig. 11C).
From this we conclude that SSY5 mRNA is not protected from NMD by the transacting factor Pub1. However, a much more thorough analysis of RBPs needs to be
conducted to sufficiently analyze the role of trans-acting factors in the protection of
81
SSY5 mRNA from NMD. We are in the early stages of developing a screen using the S.
cerevisiae RBP mutant collection and our pGAL-GFP-SSY5 constructs developed for the
translation analysis of Ssy5 (Fig. 6B).
Discussion
Here we show the novel case of the wild-type SSY5 mRNA, which has multiple
NMD targeting signals but is not degraded by NMD. Importantly, it was previously
shown that the long 3’ UTR of the SSY5 mRNA is able to target a reporter mRNA for
degradation by NMD (Obenoskey et al., 2014). Because one of the NMD-targeting
signals is able to target an mRNA from degradation by NMD we hypothesized that the
SSY5 mRNA is being protected by some mechanism from degradation by NMD.
Elucidating the mechanism that is protecting the SSY5 mRNA from NMD can add
valuable clues to understanding the role of NMD in gene regulation of wild-type mRNAs
and the potential physiologic consequences of perturbing the NMD pathway (i.e.
nonsense suppression and readthrough therapies).
Here we show that replacing the long 3’ UTR of the SSY5 mRNA with the much
shorter 3’ UTR of the ADH1 mRNA does not alter the stability of the mRNA from NMD
(Fig. 4). We see the same result upon removal of the translated SSY5 uORF (Fig. 5).
These results confirm that the mechanism of protection of the SSY5 mRNA is not in the
cis-elements of the 5’ or 3’ UTR of the SSY5 mRNA. This also rules out the role of tansacting factors that would be interfering with NMD by binding to a location within the 5’
or 3’ UTR of the SSY5 mRNA.
82
Previous studies have identified several ways in which an mRNA can be protected
from degradation by NMD. One of the ways in which an mRNA is protected from NMD
is by inhibition of translation (Carter et al., 1995; Dang et al., 2009; Low et al., 2005;
Noensie and Dietz, 2001). The trans-acting NMD factors (Upf1, Upf2 and Upf3) are
activated by a terminating ribosome, thus, the translation machinery must be active for
NMD to occur (Amrani et al., 2004; Bertram et al., 2001; Czaplinski et al., 1998;
Kisselev et al., 2003; Rospert et al., 2005). If an mRNA is not actively translated the
formation of the NMD mRNP will not be initiated. We did expect the SSY5 mRNA to be
translated due to the vital role for the Ssy5 protease in S. cerevisiae in maintaining
cellular homeostasis by initiating extracellular amino acid uptake via the activation of
AAP genes (Fig. 1). We confirmed by polyribosome analysis and fluorescence
microscopy that the SSY5 mRNA is actively translated (Fig. 6). However, endogenous
Ssy5 protein levels are quite low as we are unable to detect GFP signal when SSY5
mRNA is under the control of its native promoter.
We also tested the ideas that the SSY5 mRNA could be protected from NMD by
translational readthrough or by translation reinitiation in the 3’ UTR. The stop codon
context of the SSY5 mRNA and the unique features of the SSY5 mRNA 3’ UTR are all
conducive to either possibility (Fig. 7). However, we did not find evidence of
translational readthrough or translation reinitiation for the SSY5 mRNA within our
detection limits (Fig. 8 and Fig. 9). Further, it was previously shown that yeast mRNAs
are destabilized when there is a very low frequency of translational readthrough events on
an mRNA (≤0.5%; Keeling et al., 2004). So, if an mRNA is stabilized by translational
83
readthrough the readthrough must occur at a frequency ≥0.5%, which should yield a
detectable product in our Western analysis. In support of this, the majority of studies
providing ribosome footprint data do not indicate detectable ribosome footprints within
the SSY5 mRNA 3’ UTR (Michel et al., 2014).
We began our search for the involvement of trans-acting factors in the protection
of the SSY5 mRNA from NMD by looking at RBP Pub1. Pub1 was previously shown to
protect both GCN4 and YAP1 mRNAs from degradation by NMD (Ruiz-Echevarria and
Peltz, 2000). We show that Pub1 is not involved in the protection of the SSY5 mRNA
from NMD by looking at both steady-state accumulation and mRNA half-lives in wildtype and pub1Δ cells (Fig. 11). However, during these experiments we attempted to use
the GCN4 and YAP1 mRNAs as positive controls. In doing this we were unable to
reproduce the result that Pub1 is solely responsible for the protection of GCN4 and YAP1
mRNAs from NMD (Chapter 4). One explanation could be that we were not able to
obtain the exact strains used in the original study (Ruiz-Echevarria and Peltz, 2000). We
did, however, obtain the parent strains for the ones used in the original study and
reconstructed the strains, but were still unable to show that Pub1 definitively protects
GCN4 and YAP1 mRNAs from NMD (data not shown).
For all of our studies, we were careful to take into consideration the physiologic
role of Ssy5 when growing the cells (Fig. 1). Because Ssy5 is a critical component of the
SPS amino-acid sensing complex, we are aware that changes in the amino acid
composition of the media may cause variations in SSY5 mRNA levels and possibly in
susceptibility to NMD. However, we do not find that growing cells in rich media
84
(YAPD/YEPD) vs minimal media (amino-acid limiting) has any effect on SSY5 mRNA
levels or susceptibility to NMD (Fig. 3 and Fig. 11). Although it is important to note that
true amino-acid starvation, which would result in an inactive SPS-sensor and reduce the
need for SSY5 mRNA translation, was not able to be evaluated as the strains used in this
study require supplementation with amino acids to support the auxotrophic requirements
(Materials and Methods).
Further, in this study, we characterize a new mRNA that can be used as a control
in future NMD experiments. The ENT4 mRNA was previously identified in a screen as
an mRNA with a long 3’ UTR that is an NMD substrate. Here we show in repeated
experiments that the ENT4 mRNA is an NMD substrate, as steady-state mRNA
accumulation is increased by ≥2.0-fold in upf1Δ cells compared to wild-type cells and the
T1/2 is also ≥2.0 fold longer in upf1Δ cells compared to wild-type cells (Figs. 3A-C, Fig.
5A, Fig. 6C, Fig. 8C, and Fig. 9C). Thus, the ENT4 mRNA can serve as a positive NMD
control for wild-type gene regulation by NMD.
Here we have presented a novel case for an mRNA that should be targeted for
degradation by NMD, but is not, and the protection of the mRNA cannot be explained by
current models. Moving forward, it will be valuable to the NMD field to identify how
this mRNA is protected from NMD. We have provided the framework for narrowing
down the mechanism and now the studies can proceed in a more focused manner. We
have also developed valuable tools that can be used as we move forward. Designing a
screen for trans-acting factors that stabilize/destabilize the SSY5 mRNA will be very
informative, and we now have the GFP constructs to be able to do so. Once stabilizing
85
trans-acting factors are identified we can then determine whether they protect the SSY5
mRNA by direct or indirect binding and if the factors provide interference of the NMD
mRNP or remodeling of the NMD mRNP. This information will also help identify at
which step in the NMD pathway that protection is incurred - before substrate recognition,
or after substrate recognition but before mRNA degradation. Determining if the NMD
mRNP ever forms on the SSY5 mRNA would also provide valuable information. In
addition to looking further into trans-acting factors, it will be crucial to identify if certain
cis-elements are playing a role. Specifically, changing the start codon context,
manipulating the predicted -1 PRF sites, and changing the termination codon context
could all provide valuable results.
It is interesting to note that the human fungal pathogen Candida albicans also
possesses all of the components of the SPS sensor (Martinez and Ljungdahl, 2004).
Studies have shown that when C. albicans is unable to uptake extracellular amino acids
the fungus is not able to establish infection in a mouse model (Martinez and Ljungdahl,
2004). Further, the ability of C. albicans to establish infection requires Stp1, Ssy5 and
Ssy1 {Davis, 2011 #130. This raises the possibility that the regulation of SSY5 mRNA is
involved in pathogenicity of C. albicans. Going forward, it would be interesting to
analyze the SSY5 mRNA and its susceptibility to NMD in C. albicans.
86
Figure legends
Fig. 2-1. Ssy5p, encoded by the SSY5 mRNA is a major component of the yeast amino
acid SPS sensor. The SPS sensor is the primary complex of the yeast SPS-sensing
pathway, which is responsible for the sensing and indirect uptake of extracellular amino
acids. The SPS sensor consists of the integral membrane protein Ssy1 and two peripheral
membrane proteins Ptr3 and Ssy5. Free extracellular amino acids bind to Ssy1 inducing
a conformational change, which activates this sensing protein. When Ssy1 becomes
activated a signal is transduced via Ptr3 to the serine protease Ssy5. Ssy5 has a large Nterminal Pro-domain and catalytic C-terminal Cat-domain. These two domains are
autolytically cleaved but remain noncovalently associated as an inactive complex. When
activated, Ssy5 cleaves the N-terminal domain of the cytosolic transcription factors Stp1
and Stp2, which then enter the nucleus and activate the expression of amino acid
permease genes required to generate amino acid transporters {Ljungdahl, 2009
#109}(Conrad et al., 2014).
Fig. 2-2. SSY5 mRNA has multiple NMD-targeting signals. A) Schematic representing
the SSY5 mRNA and the approximate location of the NMD-targeting signals. B)
Ribosome profile of a region of the SSY5 mRNA 5’ leader region showing the compiled
data from all ribosome profiling studies. All three reading frames are shown. Start
codon methionines are denoted with “M” (green). () Indicates the reading frame of the
SSY5 mRNA uORF and () indicates the reading frame of the SSY5 ORF. Image was
downloaded from the GWIPS-viz browser (Michel et al., 2014).
87
Fig. 2-3. SSY5 mRNA is not degraded by NMD. A) Northern blot analysis of the steadystate accumulation of the SSY5 and ENT4 (a wild-type NMD substrate) mRNAs in wildtype (AAY277) and upf1Δ (AAY363) strains in the BY4741 background and in wild-type
(AAY187) and upf1Δ (AAY320) strains in the W303 background. Strains were grown in
rich media (YAPD). CYH2 pre-mRNA is an NMD control (CYH2 pre-mRNA is an
NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown for loading.
Fold change values are normalized and determined as a fold change ratio from wild-type;
values are an average of three independent trials. B) Northern blot analysis of the SSY5
and ENT4 mRNA half-lives in wild-type (AAY277) and upf1Δ (AAY363) strains from
the BY4741 background. Strains were grown in rich media (YAPD). 10µg/ml Thiolutin
was added to mid-log cultures at time=0 min. Half-life calculations were determined
using a graph of percent mRNA remaining at each time point during the exponential
decay. C) Northern blot analysis of the steady-state accumulation of the SSY5 and ENT4
mRNAs in wild-type (AAY277) and upf1Δ (AAY363) strains in the BY4741
background. Strains were grown in minimal media (SD + his, leu, met, ura, lys). D)
Northern blot analysis of the SSY5 and ENT4 mRNA half-lives in wild-type (AAY277)
and upf1Δ (AAY363) strains from the BY4741 background. Strains were grown in
minimal media (SD + his, leu, met, ura, lys). 10µg/ml Thiolutin was added to mid-log
cultures at time=0 min. Half-life calculations were determined using a graph of percent
mRNA remaining at each time point during the exponential decay and are the average of
three independent trials.
88
Fig. 2-4. Replacing the long 3’ UTR of SSY5 mRNA with the short 3’ UTR of the ADH1
mRNA does not destabilize the mRNA. A) Schematic of constructs. The native 475ntlong 3’ UTR of the SSY5 mRNA was replaced with the 180nt 3’ UTR of the ADH1
mRNA. B) Northern blot analysis of the steady-state accumulation of the native SSY5
mRNA in the wild-type strain (AAY277), and the SSY5+ADH1 3’ UTR mRNA in wildtype [NMD+] (AAY576) and upf1Δ (AAY601) strains in the BY4741 background. The
AGP1 mRNA is a downstream target (AAP gene) of Ssy5 activity and is shown to
confirm that manipulation of the SSY5 mRNA 3’ UTR did not influence the enzymatic
activity of the protein product. Strains were grown in rich media (YAPD). CYH2 premRNA is an NMD control (CYH2 pre-mRNA is an NMD substrate, CYH2 mRNA is not
an NMD substrate) and SCR1 is shown for loading. Fold change values are normalized
and determined as a fold change ratio from wild-type (lane 1); values are an average of
three independent trials. C) Northern blot analysis of the SSY5+ADH1 3’ UTR mRNA
half-life in wild-type [NMD+] (AAY576) and upf1Δ (AAY601) strains. Strains were
grown in rich media (YAPD). 10µg/ml Thiolutin was added to mid-log cultures at
time=0 min. Half-life calculations were determined using a graph of percent mRNA
remaining at each time point during the exponential decay.
Fig. 2-5. Disrupting the 5’ UTR of the SSY5 mRNA does not destabilize the mRNA. A)
Northern blot analysis of the steady-state accumulation of the pGAL-SSY5 mRNA in
wild-type [NMD+] (AAY630) and upf1Δ (AAY632) strains in the BY4741 background.
89
The ENT4 mRNA is a wild-type mRNA that is an NMD substrate. Strains were grown in
rich media with galactose (YAPG). CYH2 pre-mRNA is an NMD control (CYH2 premRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown
for loading. Fold change values are normalized and determined as a fold change ratio
from wild-type and are an average of three independent trials. B) Northern blot analysis
of the pGAL-SSY5 mRNA in wild-type [NMD+] (AAY630) and upf1Δ (AAY632) strains
in the BY4741 background. Strains were grown in rich media with galactose (YAPG).
10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Half-life calculations
were determined using a graph of percent mRNA remaining at each time point during the
exponential decay. C) Northern blot analysis of steady-state mRNA accumulation of the
native SSY5 mRNA in the wild-type strain (AAY277) and the pGAL-SSY5 mRNA in the
wild-type [NMD+] background (AAY630) to show the increase in accumulation as a
result of placing the SSY5 mRNA under the control of the galactose promoter. Strains
were grown in rich media with galactose (YAPG). Fold change values are normalized
and determined as a fold change ratio from wild-type. The native SSY5 mRNA appears
as a band at ~2.6 kb while the pGAL-SSY5 mRNA is slightly larger and appears as a band
at ~3.0 kb.
Fig. 2-6. SSY5 mRNA is translated. A) Polyribosome analysis of SSY5 mRNA (top) and
CYH2 pre-mRNA and mRNA (bottom) in wild-type (AAY187) and upf1Δ (AAY320)
strains. Strains were grown in YAPD. Whole cell lysate was centrifuged through a 1550% sucrose gradient. Total RNA was extracted from each fraction collected and total
90
RNA from fractions 2-19 was transferred to a membrane for Northern analysis. Fractions
corresponding to the 40S ribosomal peak are in lanes 4-5, fractions corresponding to the
60S ribosomal peak are in lanes 6-7, the fraction corresponding to the 80S ribosomal
peak is in lane 8 and polyribosome fractions are in lanes 9-15. The graph represents the
distribution of the mRNA in each fraction based on the relative corrected volume of the
mRNA present in each lane. In lanes where two bands were present only the band
corresponding to the size of the expected band for the mRNA was quantified (~2.6 kb for
SSY5 mRNA). Polysome analysis was repeated twice with similar results. B) Confocal
microscopy of haploid cells containing pGAL-GFP-SSY5 mRNA (AAY561; top) or
untagged SSY5 mRNA (AAY527; bottom). Strains are of the BY4743 background.
Cells were grown to saturation in YAPG, washed, fixed in 3.7% formaldehyde and
stained with 1.0 µg/ml Hoechst (33258) in the dark. Cells were immediately imaged.
Scale bar=5µM. C) Northern analysis of steady-state mRNA accumulation of the pGALGFP-SSY5 mRNA to confirm that there were no significant differences in accumulation
of the mRNA between wild-type background [NMD+] (AAY568; haploid of AAY561)
and upf1Δ (AAY625) strains. Strains were grown in YAPG. Fold change values are
normalized and determined as a fold change ratio from wild-type and are an average of
three independent trials.
Fig. 2-7. The SSY5 mRNA 3’ UTR is a good candidate for both stop codon readthrough
and reinitiation of translation. The 475nt sequence of the SSY5 mRNA 3’ UTR. The
ORF stop codon (TGA) is in red with a single underline. Triplet nucleotide codons are
91
highlighted in alternating gray and black corresponding to the main ORF reading frame.
The sequence underlined with a dotted blue line and terminating at the TAA stop codon
with a double red underline is the sequence that would be translated if stop codon
readthrough of the ORF TGA codon were to occur. The ATG in green is a downstream
start codon that could be used for the ribosome to reinitiate translation. Sequence in pink
that terminates with the TAG codon in red is the sequence that would be translated if
reinitiation were to occur in the SSY5 mRNA 3’ UTR.
Fig. 2-8. There is no evidence of SSY5 mRNA stop codon readthrough. A) Schematic of
the reporter constructs that were made along with a sequence snapshot to show exactly
which reading frame the GFP sequence was added in (arrows). Reporter #1 was made by
placing the GFP sequence in-frame just before the SSY5 mRNA TGA stop codon.
Reporter #2 was made by placing the GFP sequence in-frame just before the stop codon
that would be used if translational readthrough were to occur. The dark grey box
represents the SSY5 mRNA ORF while the light gray box represents the additional
sequence that would be translated should readthrough occur. B) Western blot analysis of
the Ssy5-GFP product from each strain; untagged Ssy5, lane 1 (AAY277), Ssy5-GFP
before the stop codon (reporter 1), lane 2 (AAY572), Ssy5-GFP to test for readthrough
(reporter 2), lane 3 (AAY585). Asterisks indicate where expected bands should be.
Schematics show why we see the presence of two bands (at ~104.4 kD and ~62.2 kD) for
Ssy5-GFP. The top band is unprocessed Ssy5-GFP while the bottom band is the Cterminal domain of the autolytically cleaved Ssy5-GFP. 40µg of total protein was added
92
per lane. Non-specific bands serve as an informal loading control; membranes were also
stained with Ponceau S (not shown). Western blots were done with three independent
trials and results were reproducible. C) Northern analysis of steady-state mRNA
accumulation of the SSY5, AGP1 and ENT4 mRNAs in the strains used in part (B) to
confirm that there were no significant differences in accumulation of the mRNAs
between the strains and also to confirm that addition of the GFP tag did not influence the
enzymatic activity of Ssy5. ENT4 mRNA is a wild-type mRNA that is a substrate for
NMD and AGP1 mRNA is a downstream target (AAP gene) of Ssy5. Untagged SSY5
mRNA appears as a band at ~2.6 kb while the SSY5-GFP mRNA is larger and appears as
a band at ~2.8 kb. Strains were grown in rich media (YAPD). Fold change values are
normalized and determined as a fold change ratio from wild-type (lane 1) and are an
average of three independent trials.
Fig. 2-9. There is no evidence of translation reinitiation in the SSY5 3’ UTR. A)
Schematic of the reporter constructs that were made along with a sequence snapshot to
show exactly which reading frame the GFP sequence was added in (arrows). Reporter #1
is the same as in figure 8 and was made by placing the GFP sequence in-frame just before
the SSY5 mRNA TGA stop codon. Reporter #3 was made by placing the GFP sequence
in-frame just before the stop codon that would be used if the ribosome were to reinitiate
translation in the small ORF present in the SSY5 mRNA 3’ UTR. The dark grey box
represents the SSY5 mRNA ORF while the light gray box represents the small ORF that
would be translated should the ribosome reinitiate translation. B) Western blot analysis
93
of the Ssy5-GFP product from each strain; untagged Ssy5, lane 1 (AAY277), Ssy5-GFP
to test for reinitiation (reporter 3), lane 2 (AAY581), Ssy5-GFP before the stop codon
(reporter 1), lane 3 (AAY572). Asterisks indicate where expected bands should be. Here
we only show the bottom band of the Ssy5-GFP (~62.2 kD), which is the C-terminal
domain of the autolytically cleaved Ssy5-GFP, and the bottom portion of the membrane
because the reinitiation product is very small with an expected size of ~36.6 kD. 40µg of
total protein was added per lane. Non-specific bands serve as an informal loading
control; membrane was also stained with Ponceau S (not shown). Western blots were
done with three independent trials and results were reproducible. C) Northern analysis of
steady-state mRNA accumulation of the SSY5, AGP1 and ENT4 mRNAs in the strains
used in part (B) to confirm that there were no significant differences in accumulation of
the mRNAs between the strains and also to confirm that addition of the GFP tag did not
influence the enzymatic activity of Ssy5. ENT4 mRNA is a wild-type mRNA that is a
substrate for NMD and AGP1 mRNA is a downstream target (AAP gene) of Ssy5.
Untagged SSY5 mRNA appears as a band at ~2.6 kb while the SSY5-GFP mRNA is
larger and appears as a band at ~2.8 kb. Strains were grown in rich media (YAPD). Fold
change values are normalized and determined as a fold change ratio from wild-type (lane
1) and are an average of three independent trials.
Fig. 2-10. SSY5 mRNA is translated equally efficient in wild-type and upf1Δ strains.
Western analysis of untagged Ssy5 (AAY277), and Ssy5-GFP in wild-type (AAY572)
and upf1Δ (AAY623) strains to determine if NMD has an influence on the translation
94
efficiency of the SSY5 mRNA. The construct used in this analysis was reporter #1 from
figures 8 and 9 so the expected sizes are the same (at ~104.4 kD and ~62.2 kD) and are
indicated with red asterisks. 40µg of total protein was added per lane. Non-specific
bands serve as an informal loading control; membranes were also stained with Ponceau S
(not shown). Western blots were done with three independent trials and results were
reproducible.
Fig. 2-11. SSY5 mRNA is not protected from NMD by trans-acting factor Pub1. A)
Northern blot analysis of the steady-state accumulation of SSY5 mRNA in wild-type
(AAY277), upf1Δ (AAY363), pub1Δ (AAY538), and pub1Δupf1Δ (AAY590) strains in
the BY4741 background. Strains were grown in rich media (YAPD). CYH2 pre-mRNA
is an NMD control (CYH2 pre-mRNA is an NMD substrate, CYH2 mRNA is not an
NMD substrate) and SCR1 is shown for loading. Fold change values are normalized and
determined as a fold change ratio from wild-type; values are an average of three
independent trials. B) Northern blot analysis of the steady-state accumulation of SSY5
mRNA in wild-type (AAY277), upf1Δ (AAY363), and pub1Δ (AAY538) strains in the
BY4741 background. Strains were grown in minimal media (SD + his, leu, met, ura,
lys). C) Northern blot analysis of SSY5 mRNA half-lives in wild-type (AAY277) and
pub1Δ (AAY538) strains in the BY4741 background. Strains were grown in minimal
media (SD + his, leu, met, ura, lys). 10µg/ml Thiolutin was added to mid-log cultures at
time=0 min. Half-life calculations were determined using a graph of percent mRNA
95
remaining at each time point during the exponential decay and are the average of three
independent trials.
96
Figure 2-1
97
Figure 2-2
A.
B.
98
Figure 2-3
A. Rich Media
B. Rich Media
99
Figure 2-3 (continued)
C. Minimal Media
D. Minimal Media
100
Figure 2-4
A.
B.
C.
101
Figure 2-5
A.
B.
C.
102
Figure 2-6
A.
103
Figure 2-6 (continued)
B.
C.
104
Figure 2-7
105
Figure 2-8
A.
B.
C.
106
Figure 2-9
A.
B.
C.
107
Figure 2-10
108
Figure 2-11
A. Rich Media
C. Minimal Media
B. Minimal Media
109
Table 2-1. Strains used in this study
Strain
Parent
Strain
Genotype
W303
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100
BY4741
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0
AAY320
W303
AAY363
BY4741
BY4743
AAY538
BY4741
AAY561*
BY4743
AAY568*
AAY561*
AAY572*
BY4741
AAY576*
BY4741
AAY581*
BY4741
AAY585*
BY4741
AAY590*
AAY538
AAY601*
AAY576*
AAY623*
AAY572*
AAY625*
AAY568*
AAY630*
BY4743
AAY632*
AAY630*
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100 upf1::URA3
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3
Source
Ralser et al.,
2012
Winston et al.,
1998
Atkin lab
Atkin lab
MATa/MATα his3Δ1/his3Δ1 leu2Δ0/leu2Δ0
met15Δ0/MET15 ura3Δ0/ura3Δ0 lys2Δ0/LYS2
[suc2Δ/suc2Δ]
Winston et al.,
1998
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 pub1Δ
Open
Biosystems
MATa/MATα his3Δ1/his3Δ1 leu2Δ0/leu2Δ0
met15Δ0/MET15 ura3Δ0/ura3Δ0 lys2Δ0/LYS2
[suc2Δ/suc2Δ] pGAL-GFP-HISMX6-SSY5/SSY5
MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 pGAL-GFPHISMX6-SSY5
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 GFP-HISMX6SSY5 before ORF stop codon
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SSY5
3'UTR::ADH1 3'UTR(HIS3)
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 GFP-HISMX6SSY5 before reinitiation stop codon
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 GFP-HISMX6SSY5 before readthrough stop codon
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 pub1Δ
upf1::URA3
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3 SSY5
3'UTR::ADH1 3'UTR(HIS3)
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3 GFPHISMX6-SSY5 before ORF stop codon
MATα his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3
pGAL-GFP-HISMX6-SSY5
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 pGAL-HISMX6SSY5
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3
pGAL-HISMX6-SSY5
* = new strain created for this study
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
110
References
Amrani, N., Dong, S., He, F., Ganesan, R., Ghosh, S., Kervestin, S., Li, C., Mangus,
D.A., Spatrick, P., and Jacobson, A. (2006). Aberrant termination triggers nonsensemediated mRNA decay. Biochem Soc Trans 34, 39-42.
Amrani, N., Ganesan, R., Kervestin, S., Mangus, D.A., Ghosh, S., and Jacobson, A.
(2004). A faux 3'-UTR promotes aberrant termination and triggers nonsense-mediated
mRNA decay. Nature 432, 112-118.
Andreasson, C., Heessen, S., and Ljungdahl, P.O. (2006). Regulation of transcription
factor latency by receptor-activated proteolysis. Genes Dev 20, 1563-1568.
Atkin, A.L., Altamura, N., Leeds, P., and Culbertson, M.R. (1995). The majority of yeast
UPF1 co-localizes with polyribosomes in the cytoplasm. Mol Biol Cell 6, 611-625.
Barbosa, C., Peixeiro, I., and Romao, L. (2013). Gene expression regulation by upstream
open reading frames and human disease. PLoS Genet 9, e1003529.
Bedwell, D.M., Kaenjak, A., Benos, D.J., Bebok, Z., Bubien, J.K., Hong, J., Tousson, A.,
Clancy, J.P., and Sorscher, E.J. (1997). Suppression of a CFTR premature stop mutation
in a bronchial epithelial cell line. Nat Med 3, 1280-1284.
Belew, A.T., Hepler, N.L., Jacobs, J.L., and Dinman, J.D. (2008). PRFdb: a database of
computationally predicted eukaryotic programmed -1 ribosomal frameshift signals. BMC
Genomics 9, 339.
Bertram, G., Innes, S., Minella, O., Richardson, J., and Stansfield, I. (2001). Endless
possibilities: translation termination and stop codon recognition. Microbiology 147, 255269.
Bonetti, B., Fu, L., Moon, J., and Bedwell, D.M. (1995). The efficiency of translation
termination is determined by a synergistic interplay between upstream and downstream
sequences in Saccharomyces cerevisiae. J Mol Biol 251, 334-345.
Brachmann, C.B., Davies, A., Cost, G.J., Caputo, E., Li, J., Hieter, P., and Boeke, J.D.
(1998). Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful
set of strains and plasmids for PCR-mediated gene disruption and other applications.
Yeast 14, 115-132.
Cao, D., and Parker, R. (2003). Computational modeling and experimental analysis of
nonsense-mediated decay in yeast. Cell 113, 533-545.
Carter, M.S., Doskow, J., Morris, P., Li, S., Nhim, R.P., Sandstedt, S., and Wilkinson,
M.F. (1995). A regulatory mechanism that detects premature nonsense codons in T-cell
receptor transcripts in vivo is reversed by protein synthesis inhibitors in vitro. J Biol
Chem 270, 28995-29003.
111
Chamieh, H., Ballut, L., Bonneau, F., and Le Hir, H. (2008). NMD factors UPF2 and
UPF3 bridge UPF1 to the exon junction complex and stimulate its RNA helicase activity.
Nat Struct Mol Biol 15, 85-93.
Conrad, M., Schothorst, J., Kankipati, H.N., Van Zeebroeck, G., Rubio-Texeira, M., and
Thevelein, J.M. (2014). Nutrient sensing and signaling in the yeast Saccharomyces
cerevisiae. FEMS Microbiol Rev 38, 254-299.
Cui, Y., Hagan, K.W., Zhang, S., and Peltz, S.W. (1995). Identification and
characterization of genes that are required for the accelerated degradation of mRNAs
containing a premature translational termination codon. Genes Dev 9, 423-436.
Culbertson, M.R. (1999). RNA surveillance. Unforeseen consequences for gene
expression, inherited genetic disorders and cancer. Trends Genet 15, 74-80.
Czaplinski, K., Ruiz-Echevarria, M.J., Paushkin, S.V., Han, X., Weng, Y., Perlick, H.A.,
Dietz, H.C., Ter-Avanesyan, M.D., and Peltz, S.W. (1998). The surveillance complex
interacts with the translation release factors to enhance termination and degrade aberrant
mRNAs. Genes Dev 12, 1665-1677.
Dang, Y., Low, W.K., Xu, J., Gehring, N.H., Dietz, H.C., Romo, D., and Liu, J.O.
(2009). Inhibition of nonsense-mediated mRNA decay by the natural product pateamine
A through eukaryotic initiation factor 4AIII. J Biol Chem 284, 23613-23621.
Dinman, J.D., Icho, T., and Wickner, R.B. (1991). A -1 ribosomal frameshift in a doublestranded RNA virus of yeast forms a gag-pol fusion protein. Proc Natl Acad Sci U S A
88, 174-178.
Eberle, A.B., Stalder, L., Mathys, H., Orozco, R.Z., and Muhlemann, O. (2008).
Posttranscriptional gene regulation by spatial rearrangement of the 3' untranslated region.
PLoS Biol 6, e92.
Frischmeyer, P.A., and Dietz, H.C. (1999). Nonsense-mediated mRNA decay in health
and disease. Hum Mol Genet 8, 1893-1900.
Gietz, R.D., and Woods, R.A. (2002). Transformation of yeast by lithium acetate/singlestranded carrier DNA/polyethylene glycol method. Methods Enzymol 350, 87-96.
Gonzalez, C.I., Bhattacharya, A., Wang, W., and Peltz, S.W. (2001). Nonsense-mediated
mRNA decay in Saccharomyces cerevisiae. Gene 274, 15-25.
Graber, J.H., Cantor, C.R., Mohr, S.C., and Smith, T.F. (1999). Genomic detection of
new yeast pre-mRNA 3'-end-processing signals. Nucleic Acids Res 27, 888-894.
Guan, Q., Zheng, W., Tang, S., Liu, X., Zinkel, R.A., Tsui, K.W., Yandell, B.S., and
Culbertson, M.R. (2006). Impact of nonsense-mediated mRNA decay on the global
expression profile of budding yeast. PLoS Genet 2, e203.
112
Hall, G.W., and Thein, S. (1994). Nonsense codon mutations in the terminal exon of the
beta-globin gene are not associated with a reduction in beta-mRNA accumulation: a
mechanism for the phenotype of dominant beta-thalassemia. Blood 83, 2031-2037.
Hamilton, R., Watanabe, C.K., and de Boer, H.A. (1987). Compilation and comparison of
the sequence context around the AUG startcodons in Saccharomyces cerevisiae mRNAs.
Nucleic Acids Res 15, 3581-3593.
Hammell, A.B., Taylor, R.C., Peltz, S.W., and Dinman, J.D. (1999). Identification of
putative programmed -1 ribosomal frameshift signals in large DNA databases. Genome
Res 9, 417-427.
He, F., Brown, A.H., and Jacobson, A. (1997). Upf1p, Nmd2p, and Upf3p are interacting
components of the yeast nonsense-mediated mRNA decay pathway. Mol Cell Biol 17,
1580-1594.
He, F., and Jacobson, A. (1995). Identification of a novel component of the nonsensemediated mRNA decay pathway by use of an interacting protein screen. Genes Dev 9,
437-454.
He, F., and Jacobson, A. (2001). Upf1p, Nmd2p, and Upf3p regulate the decapping and
exonucleolytic degradation of both nonsense-containing mRNAs and wild-type mRNAs.
Mol Cell Biol 21, 1515-1530.
He, F., Li, X., Spatrick, P., Casillo, R., Dong, S., and Jacobson, A. (2003). Genome-wide
analysis of mRNAs regulated by the nonsense-mediated and 5' to 3' mRNA decay
pathways in yeast. Mol Cell 12, 1439-1452.
He, F., Peltz, S.W., Donahue, J.L., Rosbash, M., and Jacobson, A. (1993). Stabilization
and ribosome association of unspliced pre-mRNAs in a yeast upf1- mutant. Proc Natl
Acad Sci U S A 90, 7034-7038.
Hentze, M.W., and Kulozik, A.E. (1999). A perfect message: RNA surveillance and
nonsense-mediated decay. Cell 96, 307-310.
Herrick, D., Parker, R., and Jacobson, A. (1990). Identification and comparison of stable
and unstable mRNAs in Saccharomyces cerevisiae. Mol Cell Biol 10, 2269-2284.
Hilleren, P., and Parker, R. (1999). Mechanisms of mRNA surveillance in eukaryotes.
Annu Rev Genet 33, 229-260.
Hogg, J.R., and Goff, S.P. (2010). Upf1 senses 3'UTR length to potentiate mRNA decay.
Cell 143, 379-389.
Hurt, J.A., Robertson, A.D., and Burge, C.B. (2013). Global analyses of UPF1 binding
and function reveal expanded scope of nonsense-mediated mRNA decay. Genome Res
23, 1636-1650.
113
Inacio, A., Silva, A.L., Pinto, J., Ji, X., Morgado, A., Almeida, F., Faustino, P., Lavinha,
J., Liebhaber, S.A., and Romao, L. (2004). Nonsense mutations in close proximity to the
initiation codon fail to trigger full nonsense-mediated mRNA decay. J Biol Chem 279,
32170-32180.
Ingolia, N.T., Ghaemmaghami, S., Newman, J.R., and Weissman, J.S. (2009). Genomewide analysis in vivo of translation with nucleotide resolution using ribosome profiling.
Science 324, 218-223.
Isken, O., and Maquat, L.E. (2008). The multiple lives of NMD factors: balancing roles
in gene and genome regulation. Nat Rev Genet 9, 699-712.
Jacobs, J.L., Belew, A.T., Rakauskaite, R., and Dinman, J.D. (2007). Identification of
functional, endogenous programmed -1 ribosomal frameshift signals in the genome of
Saccharomyces cerevisiae. Nucleic Acids Res 35, 165-174.
Kebaara, B., Nazarenus, T., Taylor, R., and Atkin, A.L. (2003). Genetic background
affects relative nonsense mRNA accumulation in wild-type and upf mutant yeast strains.
Curr Genet 43, 171-177.
Kebaara, B.W., and Atkin, A.L. (2009). Long 3'-UTRs target wild-type mRNAs for
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Nucleic Acids Res 37,
2771-2778.
Kebaara, B.W., Baker, K.E., Patefield, K.D., and Atkin, A.L. (2012). Analysis of
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Curr Protoc Cell Biol
Chapter 27, Unit 27 23.
Keeling, K.M., Lanier, J., Du, M., Salas-Marco, J., Gao, L., Kaenjak-Angeletti, A., and
Bedwell, D.M. (2004). Leaky termination at premature stop codons antagonizes
nonsense-mediated mRNA decay in S. cerevisiae. RNA 10, 691-703.
Kisselev, L., Ehrenberg, M., and Frolova, L. (2003). Termination of translation: interplay
of mRNA, rRNAs and release factors? EMBO J 22, 175-182.
Le Hir, H., Gatfield, D., Izaurralde, E., and Moore, M.J. (2001). The exon-exon junction
complex provides a binding platform for factors involved in mRNA export and nonsensemediated mRNA decay. EMBO J 20, 4987-4997.
Lee, B.S., and Culbertson, M.R. (1995). Identification of an additional gene required for
eukaryotic nonsense mRNA turnover. Proc Natl Acad Sci U S A 92, 10354-10358.
Lelivelt, M.J., and Culbertson, M.R. (1999). Yeast Upf proteins required for RNA
surveillance affect global expression of the yeast transcriptome. Mol Cell Biol 19, 67106719.
114
Lewis, B.P., Green, R.E., and Brenner, S.E. (2003). Evidence for the widespread
coupling of alternative splicing and nonsense-mediated mRNA decay in humans. Proc
Natl Acad Sci U S A 100, 189-192.
Ljungdahl, P.O. (2009). Amino-acid-induced signalling via the SPS-sensing pathway in
yeast. Biochem Soc Trans 37, 242-247.
Longtine, M.S., McKenzie, A., 3rd, Demarini, D.J., Shah, N.G., Wach, A., Brachat, A.,
Philippsen, P., and Pringle, J.R. (1998). Additional modules for versatile and economical
PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953961.
Low, W.K., Dang, Y., Schneider-Poetsch, T., Shi, Z., Choi, N.S., Merrick, W.C., Romo,
D., and Liu, J.O. (2005). Inhibition of eukaryotic translation initiation by the marine
natural product pateamine A. Mol Cell 20, 709-722.
Lykke-Andersen, J., Shu, M.D., and Steitz, J.A. (2001). Communication of the position
of exon-exon junctions to the mRNA surveillance machinery by the protein RNPS1.
Science 293, 1836-1839.
Maderazo, A.B., He, F., Mangus, D.A., and Jacobson, A. (2000). Upf1p control of
nonsense mRNA translation is regulated by Nmd2p and Upf3p. Mol Cell Biol 20, 45914603.
Maquat, L.E., and Carmichael, G.G. (2001). Quality control of mRNA function. Cell 104,
173-176.
Maquat, L.E., and Serin, G. (2001). Nonsense-mediated mRNA decay: insights into
mechanism from the cellular abundance of human Upf1, Upf2, Upf3, and Upf3X
proteins. Cold Spring Harb Symp Quant Biol 66, 313-320.
Martinez, P., and Ljungdahl, P.O. (2004). An ER packaging chaperone determines the
amino acid uptake capacity and virulence of Candida albicans. Mol Microbiol 51, 371384.
McGlincy, N.J., and Smith, C.W. (2008). Alternative splicing resulting in nonsensemediated mRNA decay: what is the meaning of nonsense? Trends Biochem Sci 33, 385393.
Mendell, J.T., Sharifi, N.A., Meyers, J.L., Martinez-Murillo, F., and Dietz, H.C. (2004).
Nonsense surveillance regulates expression of diverse classes of mammalian transcripts
and mutes genomic noise. Nat Genet 36, 1073-1078.
Michel, A.M., Fox, G., A, M.K., De Bo, C., O'Connor, P.B., Heaphy, S.M., Mullan, J.P.,
Donohue, C.A., Higgins, D.G., and Baranov, P.V. (2014). GWIPS-viz: development of a
ribo-seq genome browser. Nucleic Acids Res 42, D859-864.
115
Miyasaka, H. (1999). The positive relationship between codon usage bias and translation
initiation AUG context in Saccharomyces cerevisiae. Yeast 15, 633-637.
Muhlemann, O., Eberle, A.B., Stalder, L., and Zamudio Orozco, R. (2008). Recognition
and elimination of nonsense mRNA. Biochim Biophys Acta 1779, 538-549.
Muhlrad, D., and Parker, R. (1999). Aberrant mRNAs with extended 3' UTRs are
substrates for rapid degradation by mRNA surveillance. RNA 5, 1299-1307.
Namy, O., Hatin, I., and Rousset, J.P. (2001). Impact of the six nucleotides downstream
of the stop codon on translation termination. EMBO Rep 2, 787-793.
Neu-Yilik, G., Amthor, B., Gehring, N.H., Bahri, S., Paidassi, H., Hentze, M.W., and
Kulozik, A.E. (2011). Mechanism of escape from nonsense-mediated mRNA decay of
human beta-globin transcripts with nonsense mutations in the first exon. RNA 17, 843854.
Ni, J.Z., Grate, L., Donohue, J.P., Preston, C., Nobida, N., O'Brien, G., Shiue, L., Clark,
T.A., Blume, J.E., and Ares, M., Jr. (2007). Ultraconserved elements are associated with
homeostatic control of splicing regulators by alternative splicing and nonsense-mediated
decay. Genes Dev 21, 708-718.
Noensie, E.N., and Dietz, H.C. (2001). A strategy for disease gene identification through
nonsense-mediated mRNA decay inhibition. Nat Biotechnol 19, 434-439.
Nyiko, T., Sonkoly, B., Merai, Z., Benkovics, A.H., and Silhavy, D. (2009). Plant
upstream ORFs can trigger nonsense-mediated mRNA decay in a size-dependent manner.
Plant Mol Biol 71, 367-378.
Obenoskey, J., Lane, D.R., Atkin, A.L., and Kebaara, B.W. (2014). Immunity of the
Saccharomyces cerevisiae SSY5 mRNA to nonsense-mediated mRNA decay. Front Mol
Biosci 1, 25.
Plant, E.P., Wang, P., Jacobs, J.L., and Dinman, J.D. (2004). A programmed -1 ribosomal
frameshift signal can function as a cis-acting mRNA destabilizing element. Nucleic Acids
Res 32, 784-790.
Presnyak, V., Alhusaini, N., Chen, Y.H., Martin, S., Morris, N., Kline, N., Olson, S.,
Weinberg, D., Baker, K.E., Graveley, B.R., et al. (2015). Codon optimality is a major
determinant of mRNA stability. Cell 160, 1111-1124.
Pulak, R., and Anderson, P. (1993). mRNA surveillance by the Caenorhabditis elegans
smg genes. Genes Dev 7, 1885-1897.
Ralser, M., Kuhl, H., Ralser, M., Werber, M., Lehrach, H., Breitenbach, M., and
Timmermann, B. (2012). The Saccharomyces cerevisiae W303-K6001 cross-platform
genome sequence: insights into ancestry and physiology of a laboratory mutt. Open Biol
2, 120093.
116
Rebbapragada, I., and Lykke-Andersen, J. (2009). Execution of nonsense-mediated
mRNA decay: what defines a substrate? Curr Opin Cell Biol 21, 394-402.
Rehwinkel, J., Letunic, I., Raes, J., Bork, P., and Izaurralde, E. (2005). Nonsensemediated mRNA decay factors act in concert to regulate common mRNA targets. RNA
11, 1530-1544.
Romao, L., Inacio, A., Santos, S., Avila, M., Faustino, P., Pacheco, P., and Lavinha, J.
(2000). Nonsense mutations in the human beta-globin gene lead to unexpected levels of
cytoplasmic mRNA accumulation. Blood 96, 2895-2901.
Rospert, S., Rakwalska, M., and Dubaquie, Y. (2005). Polypeptide chain termination and
stop codon readthrough on eukaryotic ribosomes. Rev Physiol Biochem Pharmacol 155,
1-30.
Ruiz-Echevarria, M.J., and Peltz, S.W. (2000). The RNA binding protein Pub1 modulates
the stability of transcripts containing upstream open reading frames. Cell 101, 741-751.
Sharp, P.M., and Li, W.H. (1987). The codon Adaptation Index--a measure of directional
synonymous codon usage bias, and its potential applications. Nucleic Acids Res 15,
1281-1295.
Shibuya, T., Tange, T.O., Sonenberg, N., and Moore, M.J. (2004). eIF4AIII binds spliced
mRNA in the exon junction complex and is essential for nonsense-mediated decay. Nat
Struct Mol Biol 11, 346-351.
Singh, G., Jakob, S., Kleedehn, M.G., and Lykke-Andersen, J. (2007). Communication
with the exon-junction complex and activation of nonsense-mediated decay by human
Upf proteins occur in the cytoplasm. Mol Cell 27, 780-792.
Somogyi, P., Jenner, A.J., Brierley, I., and Inglis, S.C. (1993). Ribosomal pausing during
translation of an RNA pseudoknot. Mol Cell Biol 13, 6931-6940.
Tu, C., Tzeng, T.H., and Bruenn, J.A. (1992). Ribosomal movement impeded at a
pseudoknot required for frameshifting. Proc Natl Acad Sci U S A 89, 8636-8640.
Wang, W., Czaplinski, K., Rao, Y., and Peltz, S.W. (2001). The role of Upf proteins in
modulating the translation read-through of nonsense-containing transcripts. EMBO J 20,
880-890.
Welch, E.M., and Jacobson, A. (1999). An internal open reading frame triggers nonsensemediated decay of the yeast SPT10 mRNA. EMBO J 18, 6134-6145.
Williams, I., Richardson, J., Starkey, A., and Stansfield, I. (2004). Genome-wide
prediction of stop codon readthrough during translation in the yeast Saccharomyces
cerevisiae. Nucleic Acids Res 32, 6605-6616.
117
Yepiskoposyan, H., Aeschimann, F., Nilsson, D., Okoniewski, M., and Muhlemann, O.
(2011). Autoregulation of the nonsense-mediated mRNA decay pathway in human cells.
RNA 17, 2108-2118.
Zhang, S., Welch, E.M., Hogan, K., Brown, A.H., Peltz, S.W., and Jacobson, A. (1997).
Polysome-associated mRNAs are substrates for the nonsense-mediated mRNA decay
pathway in Saccharomyces cerevisiae. RNA 3, 234-244.
Zuk, D., and Jacobson, A. (1998). A single amino acid substitution in yeast eIF-5A
results in mRNA stabilization. EMBO J 17, 2914-2925.
118
CHAPTER 3
SSY5 mRNA is decay predominantly 5’ to 3’
119
Abstract
The degradation of mRNAs contributes significantly to the regulation of gene
expression as all mRNAs are subject to turnover. In Saccharomyces cerevisiae, most
cytoplasmic wild-type mRNAs are decapped in a deadenylation-dependent manner and
subsequently degraded by the 5’3’ exonuclease Xrn1. Alternatively, mRNAs can be
degraded 3’5’ by the exosome complex, albeit at a much slower rate. mRNAs that are
targeted for degradation by nonsense-mediated mRNA decay (NMD) are distinguished
from wild-type mRNAs at the early stages of mRNA decay. NMD substrates are rapidly
decapped in a deadenylation-independent manner and degraded 5’3’ by Xrn1 as well
as being rapidly deadenylated. The SSY5 mRNA is a wild-type mRNA in S. cerevisiae
with multiple NMD-targeting signals but is not degraded by NMD. We hypothesized that
investigating the genetic requirements for SSY5 mRNA decay would provide insight into
how this mRNA is degraded and when protection from NMD is incurred. Here we show
that the SSY5 mRNA is predominantly degraded 5’3’ because blocking Dcp1-mediated
decapping and Xrn1-mediated 5’3’ degradation results in increased accumulation and
stability of the SSY5 mRNA. In contrast, blocking deadenylation does not cause a
significant change in SSY5 mRNA accumulation. We also show that the 3’ end of the
SSY5 mRNA is longer in wild-type and upf1Δ strains but is shortened in strains where
decapping or 5’3’ exonucloelytic degradation is blocked suggesting that SSY5 mRNA
is deadenylated prior to 5’3’ degradation.
120
Introduction
In all organisms the fidelity of gene expression is a crucial aspect of cell health
and survival. Aberrant gene expression often result in deleterious consequences for the
organism. There are several mechanisms that act coordinately to ensure proper gene
expression. Two of these mechanisms are the opposing processes of mRNA transcription
and mRNA degradation.
In yeast, wild-type mRNAs are typically degraded in the cytoplasm by a
deadenylation-dependent mechanism that begins with shortening of the 3’ poly(A) tail to
an oligo(A) length of 10-12 nt by the Ccr4/Pop2/Not and Pan2/Pan3 deadenylase
complexes (Brown and Sachs, 1998; Decker and Parker, 1993; Tucker et al., 2002;
Tucker et al., 2001; Wahle and Winkler, 2013). Deadenylation is the rate-limiting step in
wild-type mRNA decay (Decker and Parker, 1993). Deadenylation of the mRNA
disrupts the circularization of the mRNP and exposes the 7-methylguanosine 5’ cap of the
mRNA. The coupling of translation termination to deadenylation is suggested by the
observation that translation termination factor eRF3 directly interacts poly(A) binding
protein, Pab1 (Cosson et al., 2002). Additionally, disruption of the eRF3-Pab1
interaction leads to defective deadenylation (Funakoshi et al., 2007; Kobayashi et al.,
2004). However, it has also been shown that mRNAs that never initiate translation are
also deadenylated, so translation termination does not appear to be a prerequisite for
deadenylation (Beelman and Parker, 1994; Muhlrad et al., 1995).
For wild-type mRNAs, deadenylation subsequently leads to rapid decapping of
the mRNA by the Dcp1/Dcp2 holoenzyme complex (Coller and Parker, 2004).
121
Additionally, wild-type mRNAs with poly(A) tails are resistant to decapping, and this
resistance depends on the presence of Pab1 (Beilharz and Preiss, 2007; Gallie, 1991;
Wilusz et al., 2001). Removal of the 5’ cap generates a 5’ monophosphate on the mRNA
end allowing the rapid 5’3’ mRNA degradation by the exonuclease Xrn1, which is the
predominant pathway for mRNA degradation in yeast (Decker and Parker, 1993; Hsu and
Stevens, 1993; Muhlrad et al., 1995; Muhlrad and Parker, 1994). Alternatively, mRNAs
can also be degraded 3’5’ by the exosome complex (Fig.1; Anderson and Parker, 1998;
Mangus and van Hoof, 2003; Mitchell and Tollervey, 2003).
In addition to wild-type mRNA turnover, eukaryotes have a conserved
surveillance mechanism known as the nonsense-mediated mRNA decay pathway (NMD)
that also plays a large role in the fidelity of gene expression (Bedwell et al., 1997; Conti
and Izaurralde, 2005; Hall and Thein, 1994; Hentze and Kulozik, 1999; Maquat and
Carmichael, 2001; Maquat and Serin, 2001; Pulak and Anderson, 1993). This mRNA
decay pathway is responsible for removing mRNAs that contain premature termination
codons (PTCs) from the translation pool before they lead to the buildup of truncated
proteins which could have deleterious effects (Gonzalez et al., 2001; Isken and Maquat,
2008; Muhlemann et al., 2008). Importantly, in addition to degrading transcripts with
PTCs the NMD pathway is also responsible for the regulation of a large portion of wildtype mRNAs as well. Previous work shows that ~5-20% of the yeast, Drosophila, and
human transcriptomes are affected upon inactivation of NMD (Guan et al., 2006; He et
al., 2003; Johansson et al., 2007; Lelivelt and Culbertson, 1999; Mendell et al., 2004).
Several mechanisms have been identified that target a wild-type mRNA for degradation
122
by NMD. These include: 1) a long 3’ UTR (Amrani et al., 2004; Kebaara and Atkin,
2009; Muhlrad and Parker, 1999), 2) translation of an upstream open reading frame
(uORF; Amrani et al., 2006; Barbosa et al., 2013; Guan et al., 2006; Nyiko et al., 2009),
3) a start codon in a suboptimal context which can lead to leaky scanning and out of
frame initiation of translation (Welch and Jacobson, 1999), and 4) the presence of
programmed ribosome frameshift (PRF) sites (Plant et al., 2004). The execution of NMD
for both PTC-containing substrates and wild-type substrates requires the three transacting factors Upf1, Upf2 and Upf3. Mutations or deletions in any one of the genes
encoding these factors causes significant accumulation (≥2.0 fold) of NMD substrates
(Cui et al., 1995; He et al., 1997; He and Jacobson, 1995; Lee and Culbertson, 1995;
Lelivelt and Culbertson, 1999; Maderazo et al., 2000).
Degradation of NMD substrates is different from that of wild-type mRNAs in that
NMD-substrates are rapidly decapped by Dcp1/Dcp2 without prior removal of the
poly(A) tail (Beelman et al., 1996; Cao and Parker, 2003; Hagan et al., 1995). This
deadenylation-independent decapping is likely due to improper termination of translation
in which the ribosome terminates too far from the poly(A) tail to interact with Pab1. This
is supported by the observation that tethering Pab1 close to a prematurely terminating
ribosome is able to block decapping (Coller et al., 1998). Additionally, wild-type
mRNAs with long 3’ UTRs are substrates for degradation by NMD (Amrani et al., 2004;
Kebaara and Atkin, 2009; Muhlrad and Parker, 1999), and the presence a long 3’ UTR
would mimic the context of a prematurely terminating ribosome placing the ribosome too
far from the poly(A) tail to interact with Pab1 (Amrani et al., 2004). After decapping the
123
mRNA is subjected to rapid 5’3’ degradation by Xrn1 and accelerated deadenylation
by the deadenylase complexes (Cao and Parker, 2003). However, if decapping or 5’3’
decay is blocked NMD substrates can also undergo 3’5’ decay via the exosome, but
this occurs at a much slower rate (Fig. 1; Cao and Parker, 2003; Mitchell and Tollervey,
2003; Muhlrad and Parker, 1994).
Previously, we showed that an mRNA in Saccharomyces cerevisiae, the SSY5
mRNA, has multiple NMD-targeting signals but is not degraded by the NMD pathway
(Kebaara and Atkin, 2009; Obenoskey et al., 2014; Chapter 2). We have shown that the
protection of the SSY5 mRNA from NMD is not explained by any of the known
mechanisms of protection. Thus, this mRNA is likely protected by a novel mechanism.
Determining the mechanism of protection of the SSY5 mRNA from NMD will add
valuable insight into the underlying gene regulatory mechanisms by NMD.
Since we have previously ruled out the known mechanisms of protection from
NMD for the SSY5 mRNA we have sought to determine how the SSY5 mRNA is
degraded. The comparison of SSY5 mRNA decay with decay patterns of known NMD
substrates (ENT4 mRNA and CYH2 pre-mRNA) and non-NMD substrates (PGK1 mRNA
and CYH2 mRNA) will provide clues as to how protection from NMD occurs. Here we
show that the SSY5 mRNA decay is predominately 5’3’ because mutations in the genes
encoding the enzymes required for decapping (Dcp1) and 5’3’ mRNA decay (Xrn1)
result in the accumulation and stabilization of the SSY5 mRNA. In contrast, mutations in
genes encoding components of the deadenylation complexes have no effect on SSY5
mRNA accumulation. We also show that the SSY5 mRNA is likely deadenylated prior to
124
decapping because the predominant 3’ ends of the SSY5 mRNA in dcp1Δ and xrn1Δ
strains are significantly shorter than in the wild-type and upf1Δ strains. This is the same
pattern observed for the predominant 3’ ends of the PGK1 mRNA, which is not degraded
by NMD. Alternatively, the predominant 3’ ends of the ENT4 mRNA, which is degraded
by NMD, are longer in the dcp1Δ and xrn1Δ strains than in the wild-type and upf1Δ
strains. Finally, we compare the susceptibility of the SSY5 mRNA to NMD in different
genetic backgrounds and show that protection of the SSY5 mRNA from NMD may be
background-dependent. In the commonly used BY4741 and W303 genetic backgrounds
SSY5 mRNA shows no difference in accumulation between wild-type and upf1Δ strains.
However, in a different genetic background, similar to W303, the SSY5 mRNA actually
has a FCR of 1.9±0.2 between wild-type and upf1Δ strains. This suggests that slight
changes in genetic background may influence the protection of the SSY5 mRNA from
NMD.
Materials and Methods
Yeast strains
The yeast strains used in this study are listed in Table 3-1. All yeast
transformations were done using Lithium Acetate-Mediated transformation as previously
described (Gietz and Woods, 2002). AAY589 was constructed by transforming AAY360
with the upf1Δ2 fragment from pAA70 using primers oAA48 and oAA79. AAY594,
AAY595, AAY596, AAY609, AAY610, AAY611 and AAY621 were constructed by
125
transforming the corresponding parent strain (listed in Table 3-1) with the upf1Δ6
fragment from pAA167 using primers oAA48 and oAA79.
Growth conditions
Unless otherwise noted yeast cells were grown using standard techniques with
mild agitation equivalent to 225rpm at 30˚C. All strains were grown in YAPD media,
which consists of: 1% yeast extract, 2% Bacto-Peptone, 2% dextrose, and 100mg/L
Adenine hemisulfate salt. Strains were maintained on YAPD from initial plating from
frozen stocks and all through subsequent liquid cultures.
RNA Extractions
Yeast strains were grown in 10mL cultures to an OD600 of 0.4-0.6. Cells were
harvested by centrifugation, washing in DEPC-ddH20, and flash-freezing in dry
ice/ethanol or liquid nitrogen. Cell pellets were stored at -70˚C until used for RNA
extractions. RNA extractions were performed as previously described (Kebaara et al.,
2012). RNA samples were diluted to 1µg/µl in DEPC-ddH20 and stored at -70˚C. RNA
quality check gels are performed for every RNA sample (1µl of 1µg/µl Total RNA is run
through a 0.8% agarose gel to check for degradation).
Quantitative Northern Analysis
10µg of Total RNA mixed with 3µl Formaldehyde loading dye (Ambion, cat. no.
8552) was separated through a 1.0% agarose gel containing 5.6% Formaldehyde and 1%
126
MOPS (10X MOPS: 0.2 M sodium morpholinopropanesulfonic acid (MOPS), pH 7.0,
0.05 M sodium acetate, 0.01 M EDTA; adjust pH to 7.0 with 10 M NaOH, do not
autoclave, store at room temperature in the dark.) RNA was transferred onto a
GeneScreen Plus membrane (PerkinElmer) using NorthernMax transfer buffer (Ambion,
cat. no. 8672) following the manufacturer’s protocol for downward transfer. The lane
with the RNA ladder was cut off of the gel before transfer and stained overnight in
0.5µg/ml ethidium bromide. Membranes were rinsed in 2X SSC and dried for 15
minutes at 80˚C.
Membranes were hybridized with NorthernMax prehybridization/hybridization
buffer (Ambion # 8677).
32
P-labelled probes were synthesized using ~25ng of PCR
product corresponding to the gene of interest, the RadPrime DNA Labeling System®
(Invitrogen #18428-011), and ∼50 μCi [α-32P]dCTP (3000 Ci/mmol, 10 mCi/ml) (Perkin
Elmer) following the manufacturer’s protocol. Probes are purified through a Sephadex
G-50 column equilibrated with TE pH 8.0. Membranes were hybridized overnight (12-24
hours) and then washed once at room temperature with 2X SSPE and once at 65˚C with
2X SSPE/2% SDS. Membranes were PhosphorImagedTM (GE Healthcare, Typhoon FLA
9500) and quantified using the ImageQuantTM software. All membranes were also
autoradiographed using a phosphorescent ruler to determine the size of the bands by
comparison to the RNA ladder. Membranes are stripped and stored at -20˚C for reprobing. Detailed protocol for Northern analysis can be found in Kebaara et al. (2012).
127
Poly(A) tail length analysis
This experiment was performed as described in the protocol provided with the
Poly(A) Tail-Length Analysis Kit® (Affymetrix 76455). Prior DNase treatment of RNA
samples was done using the TURBO DNA-free kit® (AM1907). Primers were designed
using the information provided in the Saccharomyces Genome Database for predicted
polyadenylation sites. Since all of the mRNAs we were interested in contained multiple
predicted poly(A) sites, the primers pairs were designed to anneal upstream of the
polyadenylation site that is predicted to be most 3’ of the other predicted sites. PCR
products were resolved on a 2.5% TAE agarose gel.
Results
SSY5 mRNA accumulation and half-life are significantly increased in xrn1Δ cells
The predominant decay route for both wild-type mRNAs and NMD substrates is
through Xrn1-mediated 5’3’ exonucloelytic degradation following removal of the 5’
mRNA cap, although this process tends to occur more rapidly for NMD substrates (Cao
and Parker, 2003). Given this, we hypothesized that the SSY5 mRNA is likely degraded
5’3’ by Xrn1, so SSY5 mRNA levels should significantly increase in an xrn1Δ strain,
but we are unsure as to what extent since we do not know the contribution of 3’5’
decay for the SSY5 mRNA. If the SSY5 mRNA accumulates to significantly high levels
(comparable to a known NMD substrate such as the ENT4 mRNA) then it is likely that
3’5’ decay rate is significantly slower.
128
When 5’3’ decay in blocked by deletion of the Xrn1 exonuclease (xrn1Δ)
quantitative Northern analysis of steady-state mRNA levels reveals a significant increase
in SSY5 mRNA accumulation compared to wild-type with a fold-change ratio (FCR) of
xrn1Δ/wild-type=7.8±0.6, n=3 (Fig. 2A). The half-life (T1/2) of the SSY5 mRNA in
xrn1Δ cells is also significantly longer at 33.9 minutes compared to 12.2 minutes in wildtype cells, n=1 (Fig. 2B). The ENT4 mRNA is shown for comparison as this mRNA is a
wild-type mRNA that is a substrate for NMD likely due to the presence of its long 3’
UTR (Kebaara and Atkin, 2009; Chapter 2). The CYH2 pre-mRNA is also shown for
comparison as a wild-type NMD substrate, whereas the CYH2 mRNA and PGK1 mRNA
are not NMD substrates (Figure 2A). We show that xrn1Δ results in a significant
increase in ENT4 mRNA accumulation indicating the 5’3’ decay mechanism is the
predominant route of decay and that exosome-mediated 3’5’ decay is not sufficient to
compensate for loss of Xrn1 (likely due to the slower rate of 3’5’ for NMD substrates;
Muhlrad and Parker, 1994). The PGK1 mRNA and CYH2 mRNA have been used
extensively in mRNA decay studies and these wild-type mRNAs are degraded by the
canonical deadenylation-dependent mRNA decay pathway (Cao and Parker, 2003;
Mitchell and Tollervey, 2003; Muhlrad and Parker, 1994; Tucker et al., 2002; Tucker et
al., 2001). Our results show that xrn1Δ has no significant impact on the accumulation of
the PGK1 mRNA indicating a much more significant role of 3’5’ decay (Fig. 2A-B;
Muhlrad and Parker, 1994). In comparison, the SSY5 mRNA yields FCRs that are much
closer to the pattern exhibited by the ENT4 and CYH2 mRNAs than the PGK1 mRNA.
These results indicate that the predominant route of degradation for SSY5 mRNA is
129
through Xrn1-mediated 5’3’ decay. This resembles the pattern for known NMD
substrates (ENT4 mRNA and CYH2 pre-mRNA) and is different than the effect of xrn1Δ
on non-NMD substrates (PGK1 and CYH2 mRNAs).
Additionally, the accumulation of SSY5 mRNA in the double mutant xrn1Δupf1Δ
is also significantly higher compared to wild-type cells with a FCR of xrn1Δupf1Δ/wildtype=9.7±0.6, n=3 (Fig. 2A). Likewise, SSY5 mRNA T1/2 in xrn1Δupf1Δ cells is
significantly longer at 40.3 minutes compared to 12.2 minutes in wild-type cells, n=1
(Fig. 2B). We do recognize that both the steady-state accumulation and T1/2 of the SSY5
mRNA in the double mutant are slightly increased from the xrn1Δ cells (FCR of
xrn1Δupf1Δ/ xrn1Δ=1.2±0.6, n=3, and difference in T1/2 of +6.4 minutes, n=1; Fig. 2AB). However, this can be accounted for by considering the slight increase in
accumulation and T1/2 of SSY5 mRNA between wild-type and upf1Δ strains alone (Fig.
2A-B). This difference is addressed in greater detail below (Fig. 6). This same pattern is
also observed for the ENT4 mRNA and CYH2 pre-mRNA.
SSY5 mRNA accumulation is modestly increased in dcp1Δ cells
During mRNA decay, Dcp1/2-mediated decapping immediately precedes 5’3’
degradation by Xrn1 (Hsu and Stevens, 1993; Muhlrad and Parker, 1994). If the mRNA
is not decapped, 5’3’ decay cannot occur because the 5’ end of the mRNA is not
exposed for exonucloelytic degradation (Stevens, 2001). Because these two events occur
in sequence, we hypothesized that blocking decapping of the mRNA by deletion of DCP1
(dcp1Δ) might yield similar results to blocking 5’3’ degradation by Xrn1 (xrn1Δ).
130
Alternatively, we might see different results because in the dcp1Δ strains the mRNAs are
capped whereas in the xrn1Δ strains the population of mRNAs includes both newly
synthesized capped mRNAs and decapped mRNAs unable to undergo 5’3’ decay (He
and Jacobson, 2001). Decapped mRNAs accumulate in p-bodies and p-bodies lack
3’5’ decay enzymes (Balagopal and Parker, 2009; Sheth and Parker, 2003). This may
lead to higher accumulation levels observed in xrn1Δ strains.
Our results show that dcp1Δ had a modest effect on the steady-state accumulation
of SSY5 mRNA with a FCR of dcp1Δ/wild-type=2.3±0.4, n=3 (Fig. 3A). This is
significantly less than the SSY5 mRNA accumulation that resulted from xrn1Δ (FCR of
dcp1Δ/wild-type=7.8±0.6, n=3; Fig. 2A). However, the T1/2 of SSY5 mRNA in dcp1Δ
cells increased to 35.9 minutes compared to 12.2 minutes in wild-type cells (n=1), which
is a similar difference to that observed with the xrn1Δ vs wild-type cells (Fig. 3B).
Similar to SSY5 mRNA, the ENT4 mRNA and CYH2 pre-mRNA also show a modest
increase in accumulation upon dcp1Δ (FCR of dcp1Δ/wild-type=1.8±0.4 and 6.4±0.5,
respectively, n=3; Fig. 3A). Interestingly, the ENT4 mRNA shows a reduction in mRNA
stability in the dcp1Δ cells compared to upf1Δ cells, although the fold change in mRNA
accumulation between the two is not quite 2-fold (Fig. 3A). The double mutant
dcp1Δupf1Δ strains also showed a different trend in accumulation between the SSY5
mRNA and the ENT4 mRNA. In contrast, accumulation of the non-NMD substrates
PGK1 and CYH2 mRNAs, as well as the half-life of the PGK1 mRNA, was the same in
wild-type, upf1Δ, dcp1Δ, and dcp1Δupf1Δ cells (Fig. 3A-B).
131
Moreover, the trend observed for the accumulation of the PGK1 mRNA is quite
similar to the trend seen in accumulation of the CYH2 mRNA in the dcp1Δ cells and
dcp1Δupf1Δ cells, which is consistent with both mRNAs being non-NMD substrates (Fig.
3A). The trend observed for the ENT4 mRNA is quite similar to the trend seen in
accumulation of the CYH2 pre-mRNA in the dcp1Δ cells and dcp1Δupf1Δ, which is
consistent with both being NMD substrates (Fig. 3A). Accumulation of the SSY5 mRNA
in the dcp1Δ cells and dcp1Δupf1Δ cells, however, does not fit the pattern of either the
CYH2 mRNA or pre-mRNA. Together these results lead us to the conclusion that SSY5
mRNA degradation is different from wild-type mRNA degradation and NMD-substrate
degradation at the step of decapping.
SSY5 mRNA accumulation is not significantly impacted by deletion of different
deadenylation components
In S. cerevisiae newly synthesized mRNAs contain poly(A) tails approximately
60-80nt in length (Beilharz and Preiss, 2007). The length of the mRNA poly(A) tail
functions, in combination with the 5’ mRNA cap, to regulate both translational efficiency
and mRNA decay (Anderson and Parker, 1998; Gallie, 1991; Muhlrad et al., 1995;
Munroe and Jacobson, 1990). The degradation of an mRNA involves a deadenylation
step regardless of the mRNA being an NMD substrate or a wild-type mRNA degraded
through the canonical pathway (Norbury, 2013). The difference is the prerequisite of
deadenylation to precede decapping for wild-type mRNA degradation where it is the ratelimiting step of mRNA decay (Cao and Parker, 2003; Decker and Parker, 1993; Muhlrad
132
and Parker, 1994). While deadenylation is not a prerequisite for decapping of NMD
substrates, the mRNAs do undergo rapid deadenylation (Cao and Parker, 2003; Muhlrad
and Parker, 1994).
Deadenylation of an mRNA occurs through the combined efforts of the
Pan2/Pan3 and the Ccr4/Pop2/Not deadenylase complexes (Wahle and Winkler, 2013;
Wiederhold and Passmore, 2010). The Pan2/Pan3 deadenylation complex was the first to
be discovered in yeast (Boeck et al., 1996; Lowell et al., 1992; Sachs and Deardorff,
1992). The Pan2 subunit is responsible for the catalytic activity of the complex (Wahle
and Winkler, 2013). The Ccr4/Pop2/Not complex is a multisubunit complex containing
at least one definitive catalytic subunit, Ccr4 (Chen et al., 2002; Tucker et al., 2002;
Tucker et al., 2001). There are conflicting reports on whether or not Caf1 is a catalytic
subunit as well in S. cerevisiae, although the catalytic activity of this subunit has been
verified in other organisms (Andersen et al., 2009; Daugeron et al., 2001; Jonstrup et al.,
2007; Liang et al., 2009; Thore et al., 2003; Tucker et al., 2002; Viswanathan et al.,
2004). It was originally shown that the Pan2/Pan3 complex acted first in the initial
trimming of the poly(A) tail in a Pab1-dependent manner (Brown and Sachs, 1998).
However, later it was observed that pan2Δ and pan3Δ strains have few defects in mRNA
deadenylation (Boeck et al., 1996; Brown et al., 1996). Together these observations
suggest that in the absence of the Pan2/Pan3 complex, the Ccr4/Pop2/Not complex is
capable of handling the full range of mRNA deadenylation (Tucker et al., 2001).
Considering these observations in the context of our previous results (Figs. 2 and 3) we
133
hypothesized that deadenylation would likely play a pertinent role in the degradation of
the SSY5 mRNA.
To test this hypothesis we analyzed the SSY5 mRNA steady-state accumulation in
several deadenylation mutant strains. Fold-change ratios between the wild-type strain
and those lacking Ccr4 (ccr4Δ strains) or Caf1 (caf1Δ strains) subunits, or both
(ccr4Δcaf1Δ strains) showed very little effect on SSY5 mRNA levels, n=3 (Fig. 4).
Likewise, deletion of the Pan2 (pan2Δ strains) catalytic subunit of the Pan2/Pan3
complex showed very little effect on SSY5 mRNA levels as well, n=3 (Fig 4). These
results are not surprising as deletion of the catalytic activity of one complex can be
compensated for by the catalytic activity of the other complex (Tucker et al., 2001).
However, the SSY5 mRNA levels showed little change in strains in which both
complexes were rendered inactive (pan2Δccr4Δ) compared to wild-type strains, n=3 (Fig.
4). A previous study showed that the MFA2 mRNA contains the longest poly(A) tail in
the pan2Δccr4Δ strain indicating a lack of deadenylation in this strain as expected.
However, decay intermediates of the MFA2 mRNA were observed in the pan2Δccr4Δ
strain indicating that at least some mRNAs in this strain can still be degraded due to a
slow rate of deadenylation-independent decapping and 5’3’ decay (Tucker et al.,
2001). Alternatively, this observation can also be explained by the possibility that the
Caf1 subunit is able to compensate for the loss of both Ccr4 and Pan2, which would
support the studies claiming Caf1 does indeed have deadenylase activity. There is also
no previous data that would support the exosome being able to compensate for loss of
deadenylase activity. Additionally, we observed the effect of NMD on the accumulation
134
of the SSY5 mRNA in the deadenylation mutant strains by deletion of Upf1 (upf1Δ
strains). We observed no significant fold changes upon Upf1 deletion in these strains
compared to strains with a wild-type Upf1 allowing us to conclude that the effect
observed for SSY5 mRNA accumulation is independent of NMD (Fig. 4). We do note
that band intensity of the upf1Δ strains does appear to be darker, however, fold change
values do not reflect a difference in SSY5 mRNA accumulation for these strains.
Originally, this was surprising, but upon closer analysis we determined this is most likely
due to the bands in the upf1Δ strains actually being thicker in width, which reflects a
slightly wider variation in mRNA lengths, but not necessarily more intense in pixel
density.
We also analyzed steady-state accumulation of NMD substrates (ENT4 mRNA
and CYH2 pre-mRNA) as well as non-NMD substrates (PGK1 and CYH2 mRNAs) for
comparison. Similar to the SSY5 mRNA, none of these mRNAs appear to be largely
effected by perturbation of deadenylation (Fig. 4). The only difference can be seen in the
upf1Δ strains for the ENT4 mRNA and CYH2 pre-mRNA, which is expected since these
are NMD substrates.
Based on these results we can conclude that blocking deadenylation as little effect
on SSY5 mRNA steady-state accumulation. However, this result is consistent with that of
both an NMD-substrate (ENT4 mRNA) and a wild-type mRNA (PGK1 mRNA).
135
SSY5 mRNA has a shortened 3’ end in xrn1Δ and dcp1Δ strains
Following the results in Figures 2-4 we wanted to determine if the SSY5 mRNA
was deadenylated and, if so, if we could determine at which step the mRNA is
deadenylated. We hypothesized that the mRNA would be deadenylated as both wild-type
mRNAs and NMD-substrates are deadenylated, but we were unsure if deadenylation was
occurring as a prerequisite to decapping. To do this we used the Poly(A) Tail-Length
Assay Kit from Affymetrix (Materials and Methods). Briefly, a poly(A) polymerase first
adds a short series of guanosine and inosine residues (G/I) to the 3’ ends of
polyadenylated mRNAs. The G/I tails become the priming site for reverse transcription
of the mRNAs to cDNA molecules. Two pairs of primers are then used to amplify the 3’
ends of target mRNAs. The first primer pair, the gene-specific (G.S.) primer pair,
consists of a forward and reverse primer that anneal to the 3’ end of the mRNA both of
which are upstream of the polyadenylation site(s). The second primer pair, the poly(A)
tail (“Tail”) primer pair, consists of the same upstream primer from the G.S. primer pair
and a universal reverse primer that anneals to the G/I tail (Fig. 5A). PCR products
obtained from the use of these two primer pairs are then compared in order to determine
the length of the 3’ poly(A) tail. It should be noted that because the poly(A) sites for the
mRNAs we used are not definitive (as there are multiple predicted sites) we designed the
G.S. primer pair to anneal upstream (3’) of all predicted poly(A) sites (SGD).
The Poly(A) Tail-Length Assay Kit was used to determine the length of the 3’
end, which includes the poly(A) tail, of steady-state SSY5, ENT4, and PGK1 mRNAs in
wild-type, upf1Δ, xrn1Δ, and dcp1Δ strains, n=2. However, an RNase H control was not
136
done so we are unable to positively confirm that the results observed are strictly due to
differences in poly(A) tail length. We can, however, use the results to analyze
differences in the 3’ end length of the mRNAs which allows us to make a preliminary
assumption about the poly(A) tail length of the mRNAs. As shown in Figure 5B, the 3’
end of the SSY5 mRNA is the same in both wild-type and upf1Δ strains (lane with the
“Tail” primers product). In contrast, the 3’ end of the SSY5 mRNA is much shorter in
both xrn1Δ and dcp1Δ strains. This indicates that in the absence of decapping or 5’3’
nucleolytic degradation the SSY5 mRNA can be shortened from the 3’ end. This same
pattern is observed for the 3’ end of the PGK1 mRNA among all four strains (Fig.5 B).
In contrast, the ENT4 mRNA shows the same 3’ end length in both wild-type and upf1Δ
strains, but in these strains the 3’ end is much shorter than in the xrn1Δ and dcp1Δ strains
(Fig.5 B). This leads to the preliminary conclusion that when the ENT4 mRNA, which is
an NMD substrate, cannot be degraded by the 5’ end then shortening of the 3’ end of the
mRNA is perturbed as well.
From these results we can conclude that the 3’ end of the SSY5 mRNA is
behaving just like wild-type mRNA (PGK1 mRNA) in contrast to an NMD substrate
(ENT4 mRNA) when comparing wild-type, upf1Δ, xrn1Δ, and dcp1Δ strains.
SSY5 mRNA has different stability in different strain backgrounds
In order to examine the behavior of the SSY5 mRNA in the decapping and 5’3’
decay mutants discussed above (Figs. 2 and 3) we utilized previously constructed strains
which are of a different genetic background (He et al., 1997; He and Jacobson, 1995). It
137
was previously shown that relative mRNA accumulation in wild-type and upf1Δ strains
can differ depending on strain background (Kebaara et al., 2003). With this knowledge
we originally analyzed SSY5 mRNA levels in both commonly used BY4741 and W303
genetic backgrounds and found no difference in susceptibility of SSY5 mRNA to NMD
(Chapter 2). However, we unexpectedly found a slight difference in the susceptibility of
the SSY5 mRNA to NMD in the decay mutants utilized in Figures 2 and 3. Figure 6
shows a direct comparison of the SSY5 mRNA steady-state accumulation in the BY4741
and W303 backgrounds along with the wild-type and upf1Δ strains from the same
background used for construction of the dcp1Δ and xrn1Δ strains (He et al., 1997; He and
Jacobson, 1995, 2001). Here we show there is no difference in SSY5 mRNA
accumulation between wild-type and upf1Δ strains in either the BY4741 or the W303
backgrounds. However, in the background used for the dcp1Δ and xrn1Δ strains, SSY5
mRNA accumulation shows a FCR of upf1Δ/wild-type =1.9±0.2, (n=3; Fig 6). Likewise,
ENT4 mRNA accumulation shows a similar FCR between wild-type and upf1Δ strains in
both the BY4741 background and the W303 background, but in the strain background
used to create the dcp1Δ and xrn1Δ strains this FCR is higher, (n=3; Fig. 6; He et al.,
1997; He and Jacobson, 1995, 2001). We created the isogenic upf1Δ strain used with this
set of strains, but the strain was constructed by transformation of the wild-type strain
(AAY360) of this genetic background with the upf1Δ2 cassette. The strain background
of this set wild-type, upf1Δ, xrn1Δ and dcp1Δ strains is published as having the same
auxotrophic markers as the W303 background that we originally tested (He et al., 1997;
He and Jacobson, 1995). However, there is clearly a difference in the accumulation of
138
the SSY5 mRNA between these strains and our original W303 strains. From these
observations we conclude that the protection of the SSY5 mRNA from NMD is not
entirely robust in all strain backgrounds. However, this can be used to our advantage in
determining what protects the SSY5 mRNA from NMD, discussed below.
Discussion
In this study we began to dissect the mechanism of SSY5 mRNA decay. The SSY5
mRNA is a wild-type mRNA with multiple NMD-targeting signals but is not degraded by
the NMD pathway (Kebaara and Atkin, 2009; Obenoskey et al., 2014; Chapter 2). We
have previously presented the case that the SSY5 mRNA is likely protected from NMD by
a novel mechanism (Chapter 2). The goal in studying the mechanism of SSY5 mRNA
decay was to determine if the mRNA was being degraded via the wild-type
deadenylation-dependent decapping pathway or by the deadenylation-independent
decapping NMD pathway. As a consequence we were hoping to be able to identify if
protection of the SSY5 mRNA from NMD is incurred before recognition as an NMD
substrate, in which we would see evidence of degradation by the deadenylationdependent decapping pathway, or if protection is incurred at a point after recognition as
an NMD substrate, in which we would potentially see evidence of deadenylationindependent decapping and decay. However, our data presented here does not provide a
clear-cut answer. Instead, we learn something quite intriguing about SSY5 mRNA decay.
When Xrn1-mediated 5’3’ decay is blocked in xrn1Δ strains, SSY5 mRNA accumulates
to significantly higher levels compared to wild-type cells (Fig. 2). However, when Dcp1-
139
mediated decapping is blocked in dcp1Δ strains, SSY5 mRNA accumulates at much lower
levels than in the xrn1Δ cells. Previous work has shown that a significant fraction of the
mRNAs in xrn1Δ cells are in fact decapped (He and Jacobson, 2001). This indicates that
the presence of the 5’ mRNA cap (in dcp1Δ cells) is able to increase the rate of exosomemediated 3’5’ degradation, while this process is much slower when the cap is removed
(in xrn1Δ cells) allowing for the greater accumulation of SSY5 mRNA (Figs. 2A and 3A).
This exact same trend is not consistent for either an NMD substrate (ENT4 mRNA) or a
wild-type mRNA (PGK1; Figs. 2A and 3A). Further, blocking deadenylation through
deletion of the catalytic subunits of either the Pan2/Pan3 deadenylase complex (pan2Δ
strains) or the Ccr4/Pop2/Not complex (ccr4Δ, caf1Δ, ccr4Δcaf1Δ strains) or both
complexes (ccr4Δpan2Δ strains) had very little effect on the steady-state accumulation of
SSY5 mRNA (Fig. 4). Together these results suggest an incredibly tight window in the
regulation of SSY5 mRNA levels as there is a significant difference in mRNA
accumulation between the dcp1Δ strains and the xrn1Δ strains, two steps that occur in
immediate sequence during 5’3’ mRNA decay (Figs. 2A and 3A).
Moreover, a previous study was done to identify direct targets of the NMD
pathway in S. cerevisiae by virtue of association with core NMD factor Upf1 (Johansson
et al., 2007). Interestingly, in this study Upf1 was found to specifically associate with the
SSY5 mRNA. An mRNA was considered to be enriched for association with Upf1 if the
signal intensity from the mRNAs co-purifying with TAP-Upf1 differed from the negative
control by at least two-fold. The SSY5 mRNA showed an enrichment of 4.76 fold over
the negative control. If enriched association of an mRNA with Upf1 is indicative of the
140
mRNA being a direct NMD substrate, as was hypothesized in this study, the SSY5 mRNA
is being recognized by the NMD machinery as an NMD substrate. According to this
interpretation, protection of the SSY5 mRNA from NMD should be incurred at a point
after substrate recognition. It is also possible that protection is incurred even earlier but I
a way that does not interfere with stable Upf1 binding.
Additionally, we found that in working with a new strain background SSY5
mRNA susceptibility to NMD was slightly different than in the BY4741 and W303
backgrounds used previously (Fig 6). Upon further contemplation, this initially
surprising and somewhat frustrating observation can actually be used to our advantage.
Previously, we determined that trans-acting factor Pub1 is not responsible for the
protection of SSY5 mRNA from NMD. We had thought this may be the case based on an
earlier study showing Pub1 is responsible for the protection of GCN4 and YAP1 mRNAs
from NMD (Ruiz-Echevarria and Peltz, 2000). This leaves us in search of potential
trans-acting factors that may be involved in the protection of SSY5 mRNA from NMD.
We have developed the strains to be able to test RNA binding protein (RBP) mutants for
the effect on SSY5 mRNA levels (Chapter 2), but taking on the entire yeast RBP
collection at once is a cumbersome task. By comparing differences between the BY4741
and W303 backgrounds with those of the dcp1Δ and xrn1Δ strain background we are
presented with a much more realistic starting point for determining which factors may be
involved in SSY5 protection from NMD.
141
Figure legends
Fig. 3-1. Wild-type mRNAs and NMD substrates are degraded by different mechanisms.
Most wild-type mRNAs are first deadenylated by the Pan2/Pan2 and Ccr4/Pop2/Not
deadenylase complexes. Shortening of the poly(A) tail leads to deadenylation-dependent
decapping by the Dcp1/Dcp2 decapping complex. Once decapped the mRNA is
subjected to 5’3’ degradation by the Xrn1 exonuclease or 3’5’ degradation by the
exosome complex. NMD substrates are rapidly decapped in a deadenylation-independent
manner, followed by rapid deadenylation and 5’3’ degradation by Xrn1. In the
absence of decapping or 5’3’ degradation NMD substrates can also undergo 3’5’
degradation, but at a much slower rate.
Fig. 3-2. SSY5 mRNA steady-state accumulation is significantly increased and it has a
significantly longer half-life in an xrn1Δ strain. A) Northern blot analysis of the steadystate accumulation of the SSY5, ENT4 (a wild-type NMD substrate), and PGK1 (a wildtype mRNA degraded by the canonical pathway) mRNAs in wild-type (AAY360), upf1Δ
(AAY589), xrn1Δ (AAY389), and xrn1Δupf1Δ (AAY611) strains grown in YAPD to
mid-log phase. CYH2 pre-mRNA is an NMD control (CYH2 pre-mRNA is an NMD
substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown for loading. Fold
change values are normalized and determined as a fold change ratio from wild-type;
values are an average of three independent trials. B) mRNA half-life analysis of the SSY5
and PGK1 mRNAs in the same strains used in part A. Strains were grown in YAPD and
10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Half-life calculations
142
were determined using a graph of percent mRNA remaining at each time point during the
exponential decay.
Fig. 3-3. SSY5 mRNA steady-state accumulation is moderately increased in a dcp1Δ
strain while half-lives are significantly stabilized in a dcp1Δ strain. A) Northern blot
analysis of the steady-state accumulation of the SSY5, ENT4, and PGK1 mRNAs in wildtype (AAY360), upf1Δ (AAY589), dcp1Δ (AAY390), and dcp1Δupf1Δ (AAY621) strains
grown in YAPD to mid-log phase. CYH2 pre-mRNA is an NMD control (CYH2 premRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown
for loading. Fold change values are normalized and determined as a fold change ratio
from wild-type; values are an average of three independent trials. B) mRNA half-life
analysis of the SSY5 and PGK1 mRNAs in the same strains used in part A. Strains were
grown in YAPD. 10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Halflife calculations were determined using a graph of percent mRNA remaining at each time
point during the exponential decay.
Fig. 3-4. Mutations altering the deadenylase complexes do not have a significant
influence on SSY5 mRNA stability. Northern blot analysis of the steady-state
accumulation of the SSY5, ENT4, and PGK1 mRNAs in wild-type (AAY391), upf1Δ
(AAY594), ccr4Δ (AAY393), and ccr4Δupf1Δ (AAY596), ccr4Δcaf1Δ (AAY394),
ccr4Δcaf1Δupf1Δ (AAY609), pan2Δ (AAY395), pan2Δupf1Δ (AAY595), pan2Δccr4Δ
(AAY396), and pan2Δccr4Δupf1Δ (AAY610) strains grown in YAPD. CYH2 pre-
143
mRNA is an NMD control (CYH2 pre-mRNA is an NMD substrate, CYH2 mRNA is not
an NMD substrate) and SCR1 is shown for loading. Fold change values are normalized
and determined as a fold change ratio from wild-type; values are an average of three
independent trials.
Fig. 3-5. SSY5 mRNA has a shortened 3’ end in xrn1Δ and dcp1Δ strains. A) Schematic
of how the Poly(A) Tail Length Analysis works. This analysis was done using the
“Poly(A) Tail-Length Assay Kit” from Affymetrix (#76455). In this assay, a G/I tail is
added to the end of mRNAs containing a Poly(A) tail. These G/I-tailed mRNAs are then
reverse transcribed to cDNA, which provides a template for subsequent PCR. PCR is
performed using two sets of primers independently. The Gene Specific (G.S.; teal) primer
pair is comprised of a forward primer that binds anywhere within the open reading frame
(ORF) or 3’ UTR and a reverse primer that binds immediately upstream (5’) of the
Poly(A) start site. *It is important to note that the exact Poly(A) start site of the mRNAs
in this figure were unknown so the reverse primer was designed just upstream of the first
predicted Poly(A) start site according to the Saccharomyces Genome Database (SGD).
The Poly(A) Tail (Tail; pink) primer pair uses the same forward primer from the G.S.
pair and a reverse primer that anneals to the G/I tail (provided with the kit). B) Products
from the PCR reactions using the above primer pairs for the SSY5, ENT4, and PGK1
mRNAs in the wild-type (AAY360), upf1Δ (AAY589), xrn1Δ (AAY389), and dcp1Δ
(AAY390) strains were resolved on a 2.5% agarose TAE gel. Strains were grown in
144
YAPD and the total RNA samples used for the reactions were the same RNA samples
that were used for Northern analysis in figures 2A and 3A.
Fig. 3-6. SSY5 mRNA has different stability in different strain backgrounds. Northern
blot analysis of the steady-state accumulation of the SSY5 and ENT4 mRNAs in wild-type
(AAY277) and upf1Δ (AAY363) in the BY4741 background, wild-type (AAY187) and
upf1Δ (AAY320) in the W303 background, and in wild-type (AAY360) and upf1Δ
(AAY589) in the “W303” background that originated from the wild-type strain in He and
Jacobson, 1995. Strains were grown in YAPD. CYH2 pre-mRNA is an NMD control
(CYH2 pre-mRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and
SCR1 is shown for loading. Fold change values are normalized and determined as a fold
change ratio from wild-type; values are an average of three independent trials.
145
Figure 3-1
146
Figure 3-2
A.
B.
147
Figure 3-3
A.
B.
148
Figure 3-4
149
Figure 3-5
A.
B.
150
Figure 3-6
151
Table 3-1. Strains used in this study
Strain
Parent
Strain
Genotype
W303
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100
BY4741
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100 upf1::URA3
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY360
100
AAY363
BY4741
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY389
AAY360
100 xrn1::ADE2
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY390
AAY360
100 dcp1::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY391
cup1Δ::LEU2PM
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY393
AAY391
cup1Δ::LEU2PM ccr4::NEO
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY394
AAY391
cup1Δ::LEU2PM ccr4::NEO caf1::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY395
AAY391
cup1Δ::LEU2PM pan2::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY396
AAY391
cup1Δ::LEU2PM ccr4::NEO pan2::URA3
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY589* AAY360
100 upf1::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY594* AAY391
cup1Δ::LEU2PM upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY595* AAY395
cup1Δ::LEU2PM pan2::URA3 upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY596* AAY393
cup1Δ::LEU2PM ccr4::NEO upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY609* AAY394
cup1Δ::LEU2PM ccr4::NEO caf1::URA3 upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY610* AAY396
cup1Δ::LEU2PM ccr4::NEO pan2::URA3 upf1::TRP1
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY611* AAY389
100 xrn1::ADE2 upf1::TRP1
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY621* AAY390
100 dcp1::URA3 upf1::TRP1
* = new strain created for this study
AAY320
W303
Source
Ralser et al.,
2012
Winston et al.,
1998
Atkin lab
He and
Jacobson, 1995
Atkin lab
He and
Jacobson, 2001
He and
Jacobson, 2001
Tucker et al.,
2001
Tucker et al.,
2001
Tucker et al.,
2001
Tucker et al.,
2001
Tucker et al.,
2001
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
152
References
Amrani, N., Dong, S., He, F., Ganesan, R., Ghosh, S., Kervestin, S., Li, C., Mangus,
D.A., Spatrick, P., and Jacobson, A. (2006). Aberrant termination triggers nonsensemediated mRNA decay. Biochem Soc Trans 34, 39-42.
Amrani, N., Ganesan, R., Kervestin, S., Mangus, D.A., Ghosh, S., and Jacobson, A.
(2004). A faux 3'-UTR promotes aberrant termination and triggers nonsense-mediated
mRNA decay. Nature 432, 112-118.
Andersen, K.R., Jonstrup, A.T., Van, L.B., and Brodersen, D.E. (2009). The activity and
selectivity of fission yeast Pop2p are affected by a high affinity for Zn2+ and Mn2+ in
the active site. RNA 15, 850-861.
Anderson, J.S., and Parker, R.P. (1998). The 3' to 5' degradation of yeast mRNAs is a
general mechanism for mRNA turnover that requires the SKI2 DEVH box protein and 3'
to 5' exonucleases of the exosome complex. EMBO J 17, 1497-1506.
Balagopal, V., and Parker, R. (2009). Polysomes, P bodies and stress granules: states and
fates of eukaryotic mRNAs. Curr Opin Cell Biol 21, 403-408.
Barbosa, C., Peixeiro, I., and Romao, L. (2013). Gene expression regulation by upstream
open reading frames and human disease. PLoS Genet 9, e1003529.
Bedwell, D.M., Kaenjak, A., Benos, D.J., Bebok, Z., Bubien, J.K., Hong, J., Tousson, A.,
Clancy, J.P., and Sorscher, E.J. (1997). Suppression of a CFTR premature stop mutation
in a bronchial epithelial cell line. Nat Med 3, 1280-1284.
Beelman, C.A., and Parker, R. (1994). Differential effects of translational inhibition in cis
and in trans on the decay of the unstable yeast MFA2 mRNA. J Biol Chem 269, 96879692.
Beelman, C.A., Stevens, A., Caponigro, G., LaGrandeur, T.E., Hatfield, L., Fortner,
D.M., and Parker, R. (1996). An essential component of the decapping enzyme required
for normal rates of mRNA turnover. Nature 382, 642-646.
Beilharz, T.H., and Preiss, T. (2007). Widespread use of poly(A) tail length control to
accentuate expression of the yeast transcriptome. RNA 13, 982-997.
Boeck, R., Tarun, S., Jr., Rieger, M., Deardorff, J.A., Muller-Auer, S., and Sachs, A.B.
(1996). The yeast Pan2 protein is required for poly(A)-binding protein-stimulated
poly(A)-nuclease activity. J Biol Chem 271, 432-438.
Brown, C.E., and Sachs, A.B. (1998). Poly(A) tail length control in Saccharomyces
cerevisiae occurs by message-specific deadenylation. Mol Cell Biol 18, 6548-6559.
Brown, C.E., Tarun, S.Z., Jr., Boeck, R., and Sachs, A.B. (1996). PAN3 encodes a
subunit of the Pab1p-dependent poly(A) nuclease in Saccharomyces cerevisiae. Mol Cell
Biol 16, 5744-5753.
153
Cao, D., and Parker, R. (2003). Computational modeling and experimental analysis of
nonsense-mediated decay in yeast. Cell 113, 533-545.
Chen, J., Chiang, Y.C., and Denis, C.L. (2002). CCR4, a 3'-5' poly(A) RNA and ssDNA
exonuclease, is the catalytic component of the cytoplasmic deadenylase. EMBO J 21,
1414-1426.
Coller, J., and Parker, R. (2004). Eukaryotic mRNA decapping. Annu Rev Biochem 73,
861-890.
Coller, J.M., Gray, N.K., and Wickens, M.P. (1998). mRNA stabilization by poly(A)
binding protein is independent of poly(A) and requires translation. Genes Dev 12, 32263235.
Conti, E., and Izaurralde, E. (2005). Nonsense-mediated mRNA decay: molecular
insights and mechanistic variations across species. Curr Opin Cell Biol 17, 316-325.
Cosson, B., Couturier, A., Chabelskaya, S., Kiktev, D., Inge-Vechtomov, S., Philippe,
M., and Zhouravleva, G. (2002). Poly(A)-binding protein acts in translation termination
via eukaryotic release factor 3 interaction and does not influence [PSI(+)] propagation.
Mol Cell Biol 22, 3301-3315.
Cui, Y., Hagan, K.W., Zhang, S., and Peltz, S.W. (1995). Identification and
characterization of genes that are required for the accelerated degradation of mRNAs
containing a premature translational termination codon. Genes Dev 9, 423-436.
Daugeron, M.C., Mauxion, F., and Seraphin, B. (2001). The yeast POP2 gene encodes a
nuclease involved in mRNA deadenylation. Nucleic Acids Res 29, 2448-2455.
Decker, C.J., and Parker, R. (1993). A turnover pathway for both stable and unstable
mRNAs in yeast: evidence for a requirement for deadenylation. Genes Dev 7, 1632-1643.
Funakoshi, Y., Doi, Y., Hosoda, N., Uchida, N., Osawa, M., Shimada, I., Tsujimoto, M.,
Suzuki, T., Katada, T., and Hoshino, S. (2007). Mechanism of mRNA deadenylation:
evidence for a molecular interplay between translation termination factor eRF3 and
mRNA deadenylases. Genes Dev 21, 3135-3148.
Gallie, D.R. (1991). The cap and poly(A) tail function synergistically to regulate mRNA
translational efficiency. Genes Dev 5, 2108-2116.
Gietz, R.D., and Woods, R.A. (2002). Transformation of yeast by lithium acetate/singlestranded carrier DNA/polyethylene glycol method. Methods Enzymol 350, 87-96.
Gonzalez, C.I., Bhattacharya, A., Wang, W., and Peltz, S.W. (2001). Nonsense-mediated
mRNA decay in Saccharomyces cerevisiae. Gene 274, 15-25.
Guan, Q., Zheng, W., Tang, S., Liu, X., Zinkel, R.A., Tsui, K.W., Yandell, B.S., and
Culbertson, M.R. (2006). Impact of nonsense-mediated mRNA decay on the global
expression profile of budding yeast. PLoS Genet 2, e203.
154
Hagan, K.W., Ruiz-Echevarria, M.J., Quan, Y., and Peltz, S.W. (1995). Characterization
of cis-acting sequences and decay intermediates involved in nonsense-mediated mRNA
turnover. Mol Cell Biol 15, 809-823.
Hall, G.W., and Thein, S. (1994). Nonsense codon mutations in the terminal exon of the
beta-globin gene are not associated with a reduction in beta-mRNA accumulation: a
mechanism for the phenotype of dominant beta-thalassemia. Blood 83, 2031-2037.
He, F., Brown, A.H., and Jacobson, A. (1997). Upf1p, Nmd2p, and Upf3p are interacting
components of the yeast nonsense-mediated mRNA decay pathway. Mol Cell Biol 17,
1580-1594.
He, F., and Jacobson, A. (1995). Identification of a novel component of the nonsensemediated mRNA decay pathway by use of an interacting protein screen. Genes Dev 9,
437-454.
He, F., and Jacobson, A. (2001). Upf1p, Nmd2p, and Upf3p regulate the decapping and
exonucleolytic degradation of both nonsense-containing mRNAs and wild-type mRNAs.
Mol Cell Biol 21, 1515-1530.
He, F., Li, X., Spatrick, P., Casillo, R., Dong, S., and Jacobson, A. (2003). Genome-wide
analysis of mRNAs regulated by the nonsense-mediated and 5' to 3' mRNA decay
pathways in yeast. Mol Cell 12, 1439-1452.
Hentze, M.W., and Kulozik, A.E. (1999). A perfect message: RNA surveillance and
nonsense-mediated decay. Cell 96, 307-310.
Hsu, C.L., and Stevens, A. (1993). Yeast cells lacking 5'-->3' exoribonuclease 1 contain
mRNA species that are poly(A) deficient and partially lack the 5' cap structure. Mol Cell
Biol 13, 4826-4835.
Isken, O., and Maquat, L.E. (2008). The multiple lives of NMD factors: balancing roles
in gene and genome regulation. Nat Rev Genet 9, 699-712.
Johansson, M.J., He, F., Spatrick, P., Li, C., and Jacobson, A. (2007). Association of
yeast Upf1p with direct substrates of the NMD pathway. Proc Natl Acad Sci U S A 104,
20872-20877.
Jonstrup, A.T., Andersen, K.R., Van, L.B., and Brodersen, D.E. (2007). The 1.4-A crystal
structure of the S. pombe Pop2p deadenylase subunit unveils the configuration of an
active enzyme. Nucleic Acids Res 35, 3153-3164.
Kebaara, B., Nazarenus, T., Taylor, R., and Atkin, A.L. (2003). Genetic background
affects relative nonsense mRNA accumulation in wild-type and upf mutant yeast strains.
Curr Genet 43, 171-177.
Kebaara, B.W., and Atkin, A.L. (2009). Long 3'-UTRs target wild-type mRNAs for
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Nucleic Acids Res 37,
2771-2778.
155
Kebaara, B.W., Baker, K.E., Patefield, K.D., and Atkin, A.L. (2012). Analysis of
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Curr Protoc Cell Biol
Chapter 27, Unit 27 23.
Kobayashi, T., Funakoshi, Y., Hoshino, S., and Katada, T. (2004). The GTP-binding
release factor eRF3 as a key mediator coupling translation termination to mRNA decay. J
Biol Chem 279, 45693-45700.
Lee, B.S., and Culbertson, M.R. (1995). Identification of an additional gene required for
eukaryotic nonsense mRNA turnover. Proc Natl Acad Sci U S A 92, 10354-10358.
Lelivelt, M.J., and Culbertson, M.R. (1999). Yeast Upf proteins required for RNA
surveillance affect global expression of the yeast transcriptome. Mol Cell Biol 19, 67106719.
Liang, W., Li, C., Liu, F., Jiang, H., Li, S., Sun, J., Wu, X., and Li, C. (2009). The
Arabidopsis homologs of CCR4-associated factor 1 show mRNA deadenylation activity
and play a role in plant defence responses. Cell Res 19, 307-316.
Lowell, J.E., Rudner, D.Z., and Sachs, A.B. (1992). 3'-UTR-dependent deadenylation by
the yeast poly(A) nuclease. Genes Dev 6, 2088-2099.
Maderazo, A.B., He, F., Mangus, D.A., and Jacobson, A. (2000). Upf1p control of
nonsense mRNA translation is regulated by Nmd2p and Upf3p. Mol Cell Biol 20, 45914603.
Mangus, D.A., and van Hoof, A. (2003). Making and breaking the message. Genome
Biol 4, 346.
Maquat, L.E., and Carmichael, G.G. (2001). Quality control of mRNA function. Cell 104,
173-176.
Maquat, L.E., and Serin, G. (2001). Nonsense-mediated mRNA decay: insights into
mechanism from the cellular abundance of human Upf1, Upf2, Upf3, and Upf3X
proteins. Cold Spring Harb Symp Quant Biol 66, 313-320.
Mendell, J.T., Sharifi, N.A., Meyers, J.L., Martinez-Murillo, F., and Dietz, H.C. (2004).
Nonsense surveillance regulates expression of diverse classes of mammalian transcripts
and mutes genomic noise. Nat Genet 36, 1073-1078.
Mitchell, P., and Tollervey, D. (2003). An NMD pathway in yeast involving accelerated
deadenylation and exosome-mediated 3'-->5' degradation. Mol Cell 11, 1405-1413.
Muhlemann, O., Eberle, A.B., Stalder, L., and Zamudio Orozco, R. (2008). Recognition
and elimination of nonsense mRNA. Biochim Biophys Acta 1779, 538-549.
Muhlrad, D., Decker, C.J., and Parker, R. (1995). Turnover mechanisms of the stable
yeast PGK1 mRNA. Mol Cell Biol 15, 2145-2156.
156
Muhlrad, D., and Parker, R. (1994). Premature translational termination triggers mRNA
decapping. Nature 370, 578-581.
Muhlrad, D., and Parker, R. (1999). Aberrant mRNAs with extended 3' UTRs are
substrates for rapid degradation by mRNA surveillance. RNA 5, 1299-1307.
Munroe, D., and Jacobson, A. (1990). Tales of poly(A): a review. Gene 91, 151-158.
Norbury, C.J. (2013). Cytoplasmic RNA: a case of the tail wagging the dog. Nat Rev Mol
Cell Biol 14, 643-653.
Nyiko, T., Sonkoly, B., Merai, Z., Benkovics, A.H., and Silhavy, D. (2009). Plant
upstream ORFs can trigger nonsense-mediated mRNA decay in a size-dependent manner.
Plant Mol Biol 71, 367-378.
Obenoskey, J., Lane, D.R., Atkin, A.L., and Kebaara, B.W. (2014). Immunity of the
Saccharomyces cerevisiae SSY5 mRNA to nonsense-mediated mRNA decay. Front Mol
Biosci 1, 25.
Plant, E.P., Wang, P., Jacobs, J.L., and Dinman, J.D. (2004). A programmed -1 ribosomal
frameshift signal can function as a cis-acting mRNA destabilizing element. Nucleic Acids
Res 32, 784-790.
Pulak, R., and Anderson, P. (1993). mRNA surveillance by the Caenorhabditis elegans
smg genes. Genes Dev 7, 1885-1897.
Ruiz-Echevarria, M.J., and Peltz, S.W. (2000). The RNA binding protein Pub1 modulates
the stability of transcripts containing upstream open reading frames. Cell 101, 741-751.
Sachs, A.B., and Deardorff, J.A. (1992). Translation initiation requires the PABdependent poly(A) ribonuclease in yeast. Cell 70, 961-973.
Sheth, U., and Parker, R. (2003). Decapping and decay of messenger RNA occur in
cytoplasmic processing bodies. Science 300, 805-808.
Stevens, A. (2001). 5'-exoribonuclease 1: Xrn1. Methods Enzymol 342, 251-259.
Thore, S., Mauxion, F., Seraphin, B., and Suck, D. (2003). X-ray structure and activity of
the yeast Pop2 protein: a nuclease subunit of the mRNA deadenylase complex. EMBO
Rep 4, 1150-1155.
Tucker, M., Staples, R.R., Valencia-Sanchez, M.A., Muhlrad, D., and Parker, R. (2002).
Ccr4p is the catalytic subunit of a Ccr4p/Pop2p/Notp mRNA deadenylase complex in
Saccharomyces cerevisiae. EMBO J 21, 1427-1436.
Tucker, M., Valencia-Sanchez, M.A., Staples, R.R., Chen, J., Denis, C.L., and Parker, R.
(2001). The transcription factor associated Ccr4 and Caf1 proteins are components of the
major cytoplasmic mRNA deadenylase in Saccharomyces cerevisiae. Cell 104, 377-386.
Viswanathan, P., Ohn, T., Chiang, Y.C., Chen, J., and Denis, C.L. (2004). Mouse CAF1
can function as a processive deadenylase/3'-5'-exonuclease in vitro but in yeast the
157
deadenylase function of CAF1 is not required for mRNA poly(A) removal. J Biol Chem
279, 23988-23995.
Wahle, E., and Winkler, G.S. (2013). RNA decay machines: deadenylation by the Ccr4not and Pan2-Pan3 complexes. Biochim Biophys Acta 1829, 561-570.
Welch, E.M., and Jacobson, A. (1999). An internal open reading frame triggers nonsensemediated decay of the yeast SPT10 mRNA. EMBO J 18, 6134-6145.
Wiederhold, K., and Passmore, L.A. (2010). Cytoplasmic deadenylation: regulation of
mRNA fate. Biochem Soc Trans 38, 1531-1536.
Wilusz, C.J., Gao, M., Jones, C.L., Wilusz, J., and Peltz, S.W. (2001). Poly(A)-binding
proteins regulate both mRNA deadenylation and decapping in yeast cytoplasmic extracts.
RNA 7, 1416-1424.
158
CHAPTER 4
The protection of the Saccharomyces cerevisiae wild-type GCN4 and YAP1 mRNAs
from degradation by NMD
159
Abstract
Nonsense-mediated mRNA decay was first discovered because of its role in the
rapid degradation of mRNAs that harbor premature termination codons (PTCs). It is now
recognized that a significant fraction of wild-type mRNAs are regulated by NMD as well.
Importantly, not all of the underlying mechanisms of the NMD pathway are understood.
Because as many as 1/3 of all genetic diseases and cancers link to NMD the development
of safe and effective therapies to treat these diseases is a pressing area of research. Given
this it is absolutely critical that we continue to elucidate the underlying molecular
mechanisms of NMD. Here we investigate two wild-type mRNAs, YAP1 and GCN4,
which both contain at least one NMD-targeting signal but neither mRNA is degraded by
NMD. Previously, it was reported that both mRNAs are protected by trans-acting factor
Pub1; however, we were unable to confirm these results. We show that both mRNAs
cofractionate with polyribosomes so are likely to be translated. We also show that
blocking 5’3’ exonucloelytic degradation results in moderate accumulation of both
mRNAs while blocking decapping has little effect on the accumulation of either mRNA.
Additionally, blocking deadenylation has little effect on the accumulation of either
mRNA. Together these results show that both YAP1 mRNA and GCN4 mRNA likely
have a unique method of degradation that is tightly regulated.
160
Introduction
The regulation of gene expression in all cells is critical for the proper functioning
of the organism. Many things contribute to the fidelity of gene expression but among the
most influential are the biogenesis of mRNA molecules through transcription and the
decay of mRNA molecules through mRNA degradation. In yeast, wild-type mRNAs are
typically degraded in the cytoplasm by a process that is initiated by removal of the 3’
poly(A) tail by the Pan2/Pan3 and Ccr4/Pop2/Not deadenylase complexes. Once the
poly(A) tail reaches a length of 10-12 adenine residues the circularization of the mRNP is
disrupted due to the lack of poly(A) binding protein (Pab1), which was originally bound
to the poly(A) tail (Brown and Sachs, 1998; Decker and Parker, 1993; Tucker et al.,
2002; Tucker et al., 2001; Wahle and Winkler, 2013). Disruption of the mRNP
circularization results in the exposure of the 5’ 7-methylguanosine to the decapping
complex Dcp1/2, which rapidly decaps the mRNA leaving a 5’ monophosphate exposed
(Coller and Parker, 2004). This 5’ monophosphate becomes the substrate for the
exoribonuclease Xrn1, which then degrades the mRNA in a 5’3’ direction. This
appears to be the primary pathway for the degradation of most yeast mRNAs (Decker and
Parker, 1993; Hsu and Stevens, 1993; Muhlrad et al., 1995; Muhlrad and Parker, 1994).
Alternatively, mRNAs can also be degraded 3’5’ by the exosome complex following
deadenylation (Anderson and Parker, 1998; Mangus et al., 2003; Mangus and van Hoof,
2003; Mitchell and Tollervey, 2003).
Another part of the fidelity of gene expression is the presence of several quality
control pathways, which are responsible for the rapid degradation of aberrant mRNA
161
transcripts (Behm-Ansmant et al., 2007; Hilleren et al., 2001; Isken and Maquat, 2007,
2008; Lykke-Andersen et al., 2001). One of these quality control pathways is the
nonsense-mediated mRNA decay pathway (NMD), which is responsible for the rapid
degradation of mRNAs that harbor premature termination codons (PTCs). If mRNAs
with PTCs were not rapidly removed from the translational pool of the cell they would
lead to the build-up of truncated proteins which could have dominant negative effects
(Gonzalez et al., 2001; Isken and Maquat, 2007; Muhlemann et al., 2008). NMD is a
conserved mRNA decay pathway which is present in all eukaryotes that have been
examined (Bedwell et al., 1997; Conti and Izaurralde, 2005; Hall and Thein, 1994;
Hentze and Kulozik, 1999; Maquat and Carmichael, 2001; Maquat and Serin, 2001;
Pulak and Anderson, 1993). Importantly, it is now recognized the NMD plays a vital role
in wild-type gene regulation as well. It has been observed that ~5-20% of the yeast,
Drosophila, and human transcriptomes are affected upon inactivation of NMD (Guan et
al., 2006; He et al., 2003; Johansson et al., 2007; Lelivelt and Culbertson, 1999; Mendell
et al., 2004). NMD requires three trans-acting factors: Upf1, Upf2 and Upf3. Mutations
or deletions in one or more of the genes encoding these factors stabilizes NMD substrates
(Cui et al., 1995; He et al., 1997; He and Jacobson, 1995; Lee and Culbertson, 1995;
Lelivelt and Culbertson, 1999; Maderazo et al., 2000).
Several mechanisms have been identified which can be responsible for targeting
wild-type mRNAs for degradation by NMD including: 1) a long 3’ UTR (Amrani et al.,
2004; Kebaara and Atkin, 2009; Muhlrad and Parker, 1999), 2) translation of an upstream
open reading frame (uORF; Amrani et al., 2006; Barbosa et al., 2013; Nyiko et al., 2009),
162
3) a start codon in a suboptimal context which can lead to leaky scanning and out of
frame initiation of translation (Welch and Jacobson, 1999), 4) the presence of
programmed ribosome frameshift (PRF) sites (Plant et al., 2004), and 5) the presence of
pre-mRNA introns and regulated alternative splicing resulting in PTCs (He et al., 1993;
Lewis et al., 2003; McGlincy and Smith, 2008; Ni et al., 2007).
The degradation of NMD substrates is different from that of most wild-type
mRNAs in that NMD substrates are rapidly decapped in a deadenylation-independent
process (Beelman et al., 1996; Cao and Parker, 2003; Hagan et al., 1995). NMD
substrates are also rapidly deadenylated but the deadenylation does not need to occur as a
prerequisite for mRNA decapping (Cao and Parker, 2003). After decapping NMD
substrates are rapidly degraded 5’3’ by Xrn1 (He and Jacobson, 2001; He et al., 2003).
NMD substrates can also be degraded 3’5’ by the exosome complex following
deadenylation, but this degradation takes place at a much slower rate (Cao and Parker,
2003; Mitchell and Tollervey, 2003; Muhlrad and Parker, 1994).
It was previously shown that two wild-type mRNAs in Saccharomyces cerevisiae, YAP1
and GCN4, which have translated uORFs, are protected from degradation by NMD
(Ruiz-Echevarria et al., 1998; Ruiz-Echevarria and Peltz, 2000; Vilela et al., 1998). In
this study we further characterize the mechanism of protection of these two wild-type
mRNAs from NMD by determining that they both co-fractionate as expected with
polyribosomes, and also by determining their stability and accumulation in various decay
and deadenylation mutants. We also show that trans-acting factor, Pub1, is not likely
responsible for the protection of these two mRNAs from NMD in contrast to what was
163
previously reported (Ruiz-Echevarria and Peltz, 2000).
Materials and Methods
Yeast strains
The yeast strains used in this study are listed in Table 4-1. All yeast
transformations were done using Lithium Acetate-Mediated transformation as previously
described (Gietz and Woods, 2002). AAY590 was constructed by transforming AAY538
with the upf1Δ2 fragment from pAA70 using primers oAA48 and oAA79. AAY589 was
constructed by transforming AAY360 with the upf1Δ2 fragment from pAA70 using
primers oAA48 and oAA79. AAY594, AAY595, AAY596, AAY609, AAY610,
AAY611 and AAY621 were constructed by transforming the corresponding parent strain
(listed in Table 4-1) with the upf1Δ6 fragment from pAA167 using primers oAA48 and
oAA79.
Growth conditions
Unless otherwise noted yeast cells were grown using standard techniques with
mild agitation equivalent to 225rpm at 30˚C. When cells are stated as being grown in a
certain type of media, the same media was used to grow plate cultures from frozen stock
and all subsequent liquid cultures. In this study, YAPD media consists of: 1% yeast
extract, 2% Bacto-Peptone, 2% dextrose, and 100mg/L Adenine hemisulfate salt; and
minimal media (SD+amino acids) consists of: 0.67% yeast nitrogen base without amino
acids, 2% dextrose, 20 mg/L L-Histidine, 30 mg/L L-Leucine, 20 mg/L L-Methionine, 20
164
mg/L Uracil, and 30 mg/L L-Lysine.
RNA Extractions
Yeast strains were grown in 10mL cultures to an OD600 of 0.4-0.6. Cells were
harvested by centrifugation, washing in DEPC-ddH20, and flash-freezing in dry
ice/ethanol or liquid nitrogen. Cell pellets were stored at -70˚C until used for RNA
extractions. RNA extractions were performed as previously described (Kebaara et al.,
2012). RNA samples were diluted to 1µg/µl in DEPC-ddH20 and stored at -70˚C. RNA
quality check gels are performed for every RNA sample (1µl of 1µg/µl Total RNA is run
through a 0.8% agarose gel to check for degradation).
Quantitative Northern Analysis
10µg of Total RNA mixed with 3µl Formaldehyde loading dye (Ambion, cat. no.
8552) was separated through a 1.0% agarose gel containing 5.6% Formaldehyde and 1%
MOPS (10X MOPS: 0.2 M sodium morpholinopropanesulfonic acid (MOPS), pH 7.0,
0.05 M sodium acetate, 0.01 M EDTA; adjust pH to 7.0 with 10 M NaOH, do not
autoclave, store at room temperature in the dark.) RNA was transferred onto a
GeneScreen Plus membrane (PerkinElmer) using NorthernMax transfer buffer (Ambion,
cat. no. 8672) following the manufacturer’s protocol for downward transfer. The lane
with the RNA ladder was cut off of the gel before transfer and stained overnight in
0.5µg/ml ethidium bromide. Membranes were rinsed in 2X SSC and dried for 15
minutes at 80˚C.
165
Membranes were hybridized with NorthernMax prehybridization/hybridization
buffer (Ambion # 8677).
32
P-labelled probes were synthesized using ~25ng of PCR
product corresponding to the gene of interest, the RadPrime DNA Labeling system
(Invitrogen #18428-011), and ∼50 μCi [α-32P]dCTP (3000 Ci/mmol, 10 mCi/ml) (Perkin
Elmer) following the manufacturer’s protocol. Probes are purified through a Sephadex
G-50 column equilibrated with TE pH 8.0. Membranes were hybridized overnight (12-24
hours) and then washed once at room temperature with 2X SSPE and once at 65˚C with
2X SSPE/2% SDS. Membranes were PhosphorImagedTM (GE Healthcare, Typhoon FLA
9500) and quantified using the ImageQuantTM software. All membranes were also
autoradiographed using a phosphorescent ruler to determine the size of the bands by
comparison to the RNA ladder. Membranes are stripped and stored at -20˚C for reprobing. Detailed protocol for Northern analysis can be found in Kebaara et al. (2012).
Polyribosome Analysis
Yeast polyribosome analysis was performed as previously described (Atkin et al.,
1995). Lysis buffer was composed of 20mM Tris-HCl pH 8.0, 140mM KCl, 1.5 mM
MgCl2, 1% Triton X, 0.1 mg/ml Cycloheximide, and 1.0 mg/ml Heparin; Cycloheximide
and Heparin were made fresh and added just before use. Gradient buffer (50% and 15%
sucrose) was composed of 20mM Tris-HCl pH 8.0, 140mM KCl, 5.0 mM MgCl2, 0.1
mg/ml Cycloheximide, 0.5 mg/ml Heparin, and 0.5mM DTT; Cycloheximide, Heparin,
and DTT were made fresh and added just before use. 15%-50% sucrose gradients were
made by hand, frozen at -70˚C and thawed at 4˚C overnight just before use. After
166
fractionation RNA was extracted from each fraction using acid phenol/chloroform
extraction. Total RNA from each fraction collected was loaded onto an RNA Northern
gel and transferred to a GeneScreen Plus membrane as described above. Membranes
were hybridized with a 32P-labelled probe as described above.
Poly(A) tail length analysis
This experiment was performed as described in the protocol provided with the
Poly(A) Tail-Length Analysis Kit® (Affymetrix 76455). Prior DNase treatment of RNA
samples was done using the TURBO DNA-free kit® (AM1907). Primers were designed
using the information provided in the Saccharomyces Genome Database for predicted
polyadenylation sites. Since all of the mRNAs we were interested in contained multiple
predicted poly(A) sites, the primers pairs were designed to anneal upstream of the
polyadenylation site that is predicted to be most 3’ of the other predicted sites. PCR
products were resolved on a 2.5% TAE agarose gel.
Results
YAP1 mRNA is protected from degradation by NMD while GCN4 mRNA is
partially protected from degradation by NMD
A previous study showed that the uORF-containing GCN4 and YAP1 mRNAs are
not degraded by the NMD pathway despite presence of an NMD-targeting signal (RuizEchevarria et al., 1998). Because we recently identified the SSY5 mRNA as a wild-type
mRNA that is also protected from NMD we were hoping to include the GCN4 and YAP1
167
mRNAs in our studies as well (Kebaara and Atkin, 2009; Chapter 2).
In order to validate these results we grew S.cerevisiae cells in rich growth medium
(YAPD) and confirmed by quantitative Northern analysis that steady-state accumulation
of YAP1 and GCN4 mRNAs show no significant fold change (FC) between wild-type and
upf1Δ strains. Additionally, because it has been shown that different genetic
backgrounds of yeast strains can result in varying sensitivity of mRNAs to NMD we
tested the steady-state accumulation of these mRNAs in two of the most commonly used
genetic backgrounds, BY4741 and W303 (Kebaara et al., 2003). We found that the YAP1
mRNA is protected from significant degradation by NMD in both the BY4741 and W303
genetic backgrounds with a FCR of upf1Δ/wild-type=1.4±0.1 and 1.3±0.2, respectively
(Fig. 1A). Interestingly, in the upf1Δ strains we consistently notice a slower migrating
band for the YAP1 mRNA, which appears at a much lower intensity (Fig. 1A). For the
GCN4 mRNA we do not see ≥2.0 fold change in accumulation between wild-type and
upf1Δ strains that would be indicative of an NMD substrate. However, the stead-state
accumulation of the GCN4 mRNA is consistently elevated in the upf1Δ strains in both the
BY4741 and W303 genetic backgrounds with a FCR of upf1Δ/wild-type =1.9±0.1 and
1.6±0.1, respectively (Fig. 1A). The ENT4 mRNA is shown because it is a wild-type
mRNA in S. cerevisiae that has a long 3’ UTR and is an NMD substrate (Chapter 2).
Additionally, the CYH2 pre-mRNA is an NMD substrate while the mature CYH2 mRNA
is not. SCR1 is shown for the loading control.
Further, in rich media the half-life (T1/2) of the YAP1 mRNA exceeds 60 minutes
in both the wild-type and upf1Δ strains (Fig. 1B). The T1/2 of the GCN4 mRNA is also
168
not significantly different between wild-type (T1/2=47.0min) and upf1Δ strains
(T1/2=47.5min; Fig. 1B). Based on the T1/2 analysis of the YAP1 and GCN4 mRNAs,
neither mRNA appears to be a substrate for NMD.
The YAP1 and GCN4 mRNAs do not appear to be substrates for the NMD
pathway when cells are grown in rich media (Fig. 1A-B). However, the GCN4 mRNA
codes for a transcriptional activator of amino acid biosynthetic genes, thus we
hypothesized that the GCN4 mRNA might have varying stability in different growth
media that had decreased availability of extracellular amino acids. To test this hypothesis
we grew the cells in minimal media which contained only the amino acids necessary to
support the auxotrophies of the strains (Materials and Methods). Consistent with results
in rich media, the steady-state accumulation of the GCN4 mRNA was only moderately
increased and not significantly different between wild-type and upf1Δ strains
(FCR=1.5±0.3; Fig. 1C). Additionally, the T1/2 of the GCN4 mRNA was not
significantly different between wild-type (T1/2=21.2±3.7 min) and upf1Δ strains
(T1/2=20.7±1.0 min) when cells were grown in minimal media (Fig. 1D). However, the
T1/2 in both the wild-type and upf1Δ strains is much longer when the cells are grown in
rich media compared to when the cells are grown in minimal media (Figs. 1B and 1D).
So, the difference in growth media does influence GCN4 mRNA stability but not as a
result of NMD. This same pattern was observed for the SSY5 mRNA in Chapter 2.
In yeast, the YAP1 mRNA codes for a transcription factor required for oxidative stress
tolerance. However, since we already had the membranes prepared we also analyzed the
YAP1 mRNA when cells were grown in minimal media. Consistent with the results from
169
cells grown in rich media, the YAP1 mRNA steady-state accumulation was not
significantly different between wild-type and upf1Δ strains when cells were grown in
minimal media (FCR of upf1Δ/wild-type=1.1±0.2; Fig. 1C). Likewise, the T1/2 of the
YAP1 mRNA was not significantly different between wild-type (T1/2=15.3±1.5 min) and
upf1Δ strains (T1/2=14.0±0.9 min) when cells were grown in minimal media (Fig. 1D).
But again, just like the GCN4 and SSY5 mRNAs, the T1/2 for the YAP1 mRNA in both the
wild-type and upf1Δ strains is much longer when the cells are grown in rich media
compared to when the cells are grown in minimal media (Figs. 1B and 1D).
Together, these results allow us to confirm that both the GCN4 and YAP1 mRNAs
are wild-type mRNAs that are not likely substrates for the NMD pathway. However, the
regulation of the GCN4 mRNA is not quite as clear as previously shown (RuizEchevarria et al., 1998; Ruiz-Echevarria and Peltz, 2000; Vilela et al., 1998).
YAP1 and GCN4 mRNAs are not protected from NMD by trans-acting factor Pub1
A follow up study to the one that originally showed YAP1 and GCN4 mRNAs
were protected from NMD identified poly(U) binding protein, Pub1, as being the transacting factor responsible for this protection from NMD (Ruiz-Echevarria and Peltz,
2000). In light of our previous work which identified the SSY5 mRNA as a wild-type
mRNA that is protected from NMD these results piqued our interest (Kebaara and Atkin,
2009; Chapter 2). We hypothesized that if Pub1p was involved in protection of the YAP1
and GCN4 mRNAs from NMD then it could also be involved in SSY5 mRNA protection
from NMD. We did analyze SSY5 mRNA steady-state accumulation in wild-type, upf1Δ,
170
pub1Δ, and upf1Δpub1Δ strains and did not find any influence of Pub1 on SSY5 mRNA
susceptibility to NMD (Chapter 2). However, during this analysis the YAP1 and GCN4
mRNAs were included as positive controls since it was previously shown that Pub1 does
protect these mRNAs from NMD (Ruiz-Echevarria and Peltz, 2000). In contrast to the
previous study we do not show a Pub1-dependent protection of the YAP1 and GCN4
mRNAs from NMD (Fig. 2).
When cells are grown in either rich or minimal media we do not see a significant
increase in YAP1 mRNA stability upon deletion of Pub1 with a FCR of pub1Δ/wildtype=0.9±0.2 in rich media and a FCR of pub1Δ/wild-type=0.9±0.0 in minimal media
(Fig. 1A-B). Likewise, we do not see a significant increase in GCN4 mRNA stability
upon deletion of Pub1 with a FCR of pub1Δ/wild-type=0.8±0.0 in rich media and a FCR
of pub1Δ/wild-type=0.8±0.1 in minimal media (Fig. 1A-B). Further, T1/2 analysis in
pub1Δ strains showed no significant difference in T1/2 of the YAP1 mRNA in wild-type
(T1/2=13.2±0.4 min) versus pub1Δ strains (T1/2=16.3±2.5 min; Fig. 2C). The GCN4
mRNA also showed no difference in T1/2 between wild-type (T1/2=20.5±5.1 min) and
pub1Δ strains (T1/2=24.1±4.2 min; Fig. 2C).
It should be noted that we were unable to obtain the exact strains used in the
original publication as the lab is no longer in operation (Ruiz-Echevarria and Peltz,
2000). However, we analyzed steady-state mRNA accumulation in multiple strain
backgrounds including the parent strains for those used in the publication and were still
unable to see any effect of pub1Δ on mRNA stability (data not shown).
From this we conclude that trans-acting factor Pub1 is not responsible for YAP1 or GCN4
171
mRNA protection from NMD in the strain backgrounds and conditions that we used.
Thus, there is likely another mechanism of protection that is stabilizing these two
mRNAs.
YAP1 and GCN4 mRNAs co-fractionate with polyribosomes
Since we have confirmed that the YAP1 and GCN4 mRNAs are wild-type mRNAs
that are not degraded by NMD (Fig. 1), and we are unable to confirm that trans-acting
factor Pub1 is responsible for this protection (Fig. 2), we wanted to try to identify the
mechanism of protection of these two mRNAs. One way in which mRNAs can be
protected from NMD is by inhibition of translation (Oliveira and McCarthy, 1995;
Shoemaker and Green, 2012). We hypothesize that both YAP1 and GCN4 mRNAs are
translated because of the vital functions of their protein products within the cell.
To confirm whether or not the YAP1 and GCN4 mRNAs are likely translated we
performed a polyribosome analysis for each of these mRNAs. As expected, the YAP1
mRNA is present in the polyribosome fractions (Fig. 3A). There is no significant
difference in the YAP1 mRNA co-fractionation with polyribosomes between wild-type
and upf1Δ strains other than the presence of the larger migrating band in the upf1Δ strain,
which is also consistently present in the upf1Δ strain in the steady-state and T1/2 analysis
(Fig. 3A; Fig. 1A-B). Interestingly, this slower migrating band of the YAP1 mRNA
behaves just like the pre-mRNA of CYH2, which is an NMD substrate, in the
polyribosome analysis with the larger migrating band accumulating to higher levels in the
lighter fractions in the upf1Δ strain (Chapter 2, Fig. 6). The pre-mRNA of CYH2
172
contains an intron, but the sequence analysis for YAP1 mRNA does not indicate the
presence of an intron, which would generate a pre-mRNA (SGD). This larger migrating
band could be a longer form of the mRNA generated by translation initiation at an
alternate upstream AUG, the one of the uORF for instance. In this case the ribosome
would translate a significant portion of the 5’ leader of the YAP1 mRNA and proceed to
translate in an alternate reading frame which ultimately results in the introduction of a
PTC. This would explain why the longer mRNA is a substrate for NMD and is not
detected in wild-type strains. Alternatively, the ribosome may readthrough the ORF stop
codon and terminate downstream leading to the generation of a longer mRNA, however,
this does not readily explain the sensitivity of the larger migrating band to NMD.
The GCN4 mRNA also associates with polyribosomes as expected. The GCN4
mRNA contains four short uORFs preceding the ORF. Translation of these uORFs and
the scanning of the small ribosomal subunit along the 5’ leader is what controls
translation of the GCN4 ORF where translation of the uORFs generally represses GCN4
ORF translation (Grant and Hinnebusch, 1994; Gunisova et al., 2016; Hinnebusch, 2005).
Because these short uORFs are continually translated more actively than the GCN4 ORF
it is expected that the GCN4 mRNA will co-fractionate in the lighter fractions as the
mRNA is associated with fewer ribosomes which occupy the short (2-3 codons) uORFs
(Hinnebusch, 2005). Consistent with this, we do see the GCN4 mRNA heavily present in
the lighter fractions and more faintly present in the heavier fractions (Fig. 3B). Thus,
although the GCN4 ORF may not be actively translated at all times, the mRNA is still cofractionating in a manner indicating that the mRNA is associating with ribosomes.
173
Importantly, the polyribosome profile of the GCN4 mRNA is not significantly different
between wild-type and upf1Δ strains (Fig. 3B).
From this we can conclude that both YAP1 and GCN4 mRNAs are likely to
associate with polyribosomes as they co-fractionate in a pattern that is indicative of
polyribosome association. However, further studies are needed to confirm the production
of Yap1 and Gcn4 protein products.
YAP1 and GCN4 mRNAs accumulate significantly in xrn1Δ strains
The predominant decay route for both wild-type mRNAs and NMD substrates is
through Xrn1-mediated 5’3’ exonucloelytic degradation, although this process tends to
occur more rapidly for NMD substrates (Cao and Parker, 2003). Given this, we
hypothesized that the YAP1 and GCN4 mRNAs are likely degraded 5’3’ by Xrn1.
However, the levels to which these mRNAs accumulate in an xrn1Δ strain will give us an
indication of the contribution of 3’5’ mRNA decay by the exosome.
Upon deletion of the cytoplasmic 5’3’ exonuclease, Xrn1, both YAP1 and GCN4
mRNAs accumulate to significantly higher levels compared to wild-type strains. The
YAP1 mRNA had a FCR of xrn1Δ/wild-type=6.7±0.9 and GCN4 mRNA had a FCR of
xrn1Δ/wild-type=3.8±0.3 (Fig. 4A). It is interesting to note that though the fold changes
are significantly increased in xrn1Δ strain for the YAP1 and GCN4 mRNAs accumulation
is not nearly as high as what is seen for the ENT4 mRNA (a wild-type NMD substrate),
but also differ from the PGK1 mRNA (a wild-type mRNA degraded by the
deadenylation-dependent decapping pathway) which shows no significant difference in
174
fold change (Fig. 4A). Additionally, the fold changes for the YAP1 and GCN4 mRNAs in
the xrn1Δ strain are less than what was seen for the SSY5 mRNA (Chapter 2).
Consistently, the T1/2 analysis of the YAP1 and GCN4 mRNAs both indicate
significant stabilization of the mRNAs in the xrn1Δ strain (Fig. 4B). The YAP1 mRNA
has a T1/2 of 19.2 minutes in wild-type strains compared to a T1/2 of 31.3 minutes in the
xrn1Δ strain. Likewise, the GCN4 mRNA has a T1/2 of 28.1 minutes in wild-type strains
compared to a T1/2 of 44.9 minutes in the xrn1Δ strain.
From this we can conclude that 5’3’ exonucloelytic degradation by Xrn1 is a
significant contributor to the degradation of both the YAP1 and GCN4 mRNAs.
However, because the steady-state mRNA accumulation fold changes in the xrn1Δ strain
are not quite as significant as for other mRNAs we have looked at (e.g. SSY5 and ENT4)
the 3’5’ decay is likely more efficient for the YAP1 and GCN4 mRNAs.
YAP1 and GCN4 mRNAs do not accumulate in dcp1Δ strains, but half-life analysis
shows that dcp1Δ stabilizes the mRNAs albeit to different extents
When degradation of an mRNA is initiated the Dcp1/2 decapping complex
removes the 5’ 7-methylguanosine cap to expose a 5’ monophosphate which becomes the
immediate substrate of the Xrn1 5’3’ exonuclease (Coller and Parker, 2004). If the
mRNA is not decapped, 5’3’ decay cannot occur because the 5’ end of the mRNA is
not exposed for exonucloelytic degradation by Xrn1 (Decker and Parker, 1993; Hsu and
Stevens, 1993; Muhlrad et al., 1995; Muhlrad and Parker, 1994). Given this, we
hypothesized that blocking decapping of the mRNA by deletion of Dcp1 (dcp1Δ) would
175
yield similar results to blocking 5’3’ degradation by Xrn1 (xrn1Δ) as these two events
occur in immediate sequence.
Surprisingly, when the mRNA decapping complex Dcp1/2 is rendered inactive by
deletion of the Dcp1 catalytic subunit the steady-state accumulations of the YAP1 and
GCN4 mRNAs are not significantly affected compared to the wild-type strain. The YAP1
mRNA only shows a FCR of dcp1Δ/wild-type=1.7±0.1, which is equivalent to the
accumulation seen in the upf1Δ strain (FCR of dcp1Δ/wild-type=1.8±0.0) and is not
considered to be significant (Fig. 5A). Similarly, the GCN4 mRNA shows a FCR of
dcp1Δ/wild-type=1.2±0.0, which is even less than the accumulation seen in the upf1Δ
strain (FCR of dcp1Δ/wild-type=2.0±0.2; Fig. 5A). Both the YAP1 and GCN4 mRNAs
exhibit a pattern that is similar to the wild-type NMD substrate ENT4 mRNA.
However, the T1/2 analysis in these strains reveals a bit of a different story,
especially for the YAP1 mRNA. Compared to the wild-type strain, in which the YAP1
mRNA has a T1/2=19.2 minutes, the T1/2 of the YAP1 mRNA in the dcp1Δ strain is more
than double that in the wild-type strain at 51.2 minutes (Fig 5B). For the GCN4 mRNA
the difference is not quite as significant but is still increased compared to the steady-state
mRNA accumulations. In the wild-type strain the GCN4 mRNA has a T1/2=28.1 minutes
compared to 39.2 minutes in the dcp1Δ strain (Fig 5B).
Although mRNA stability measurements are typically more indicative of what is
actually happening to the mRNA, the trend seen in the mRNA steady-state accumulations
for the YAP1 and GCN4 mRNAs cannot simply be ignored. From these results we can
conclude that at steady-state levels in the cell the YAP1 and GCN4 mRNAs are able to be
176
efficiently degraded 3’5’ by the exosome when decapping is blocked. When
considering the T1/2 analysis of the YAP1 mRNA, blocking of decapping actually
stabilizes the mRNA even more than in the xrn1Δ strain (Fig. 4B and Fig. 5B). The same
is not true for the GCN4 mRNA, however, where blocking decapping has a relatively
similar effect to deletion of Xrn1 (Fig. 4B and Fig. 5B). Up until now our analysis has
shown similar behavior of the YAP1 and GCN4 mRNAs. However, the decay of these
two mRNAs seems to be slightly different in the influence of decapping of the mRNAs.
Some strains show greater YAP1 and GCN4 mRNA susceptibility to NMD
As was noticed with the SSY5 mRNA, the YAP1 and GCN4 mRNAs appear to
have differential susceptibility to NMD in the different strain backgrounds used in these
studies. We originally only tested the susceptibility of the YAP1 and GCN4 mRNAs to
NMD in the commonly used BY4741 and W303 backgrounds. Using these strains we
saw no difference in YAP1 or GCN4 mRNA susceptibility to NMD (Fig. 1). However,
when we began using the dcp1Δ and xrn1Δ strains we noticed a difference in both YAP1
and GCN4 mRNA accumulation in the wild-type versus upf1Δ strains from the same
background (Figs. 4A and 5A). The YAP1 and GCN4 mRNAs show a steady-state FCR
of upf1Δ/wild-type=1.8±0.0 and 2.0±0.2, respectively (Figs. 4A and 5A). Likewise,
ENT4 mRNA accumulation shows a similar FCR between wild-type and upf1Δ strains in
both the BY4741 background and the W303 background, but in the strain background
used to create the dcp1Δ and xrn1Δ strains this FCR is increased (compare ENT4 mRNA
in Fig. 1A with ENT4 mRNA in Figs. 4A and 5A). The wild-type and upf1Δ strains from
177
the same background as the xrn1Δ and dcp1Δ strains were genetically constructed in the
W303 background and the auxotrophic markers are published as being the same as the
W303 strains that we originally tested (He et al., 1997; He and Jacobson, 1995, 2001).
However, there is clearly a difference in the accumulation of the YAP1 and GCN4
mRNAs between these strains and our original W303 strains. Identifying the
difference(s) between the strains from our earlier studies and the strains showing
increased NMD susceptibility will be very beneficial in determining how the YAP1 and
GCN4 mRNAs are protected from NMD.
Deadenylation mutants show little effect on YAP1 and GCN4 mRNA stability
The length of the mRNA poly(A) tail (~60-80 nt at steady state in S. cerevisiae)
functions, in combination with the 5’ mRNA cap, to regulate both the translational
efficiency of the mRNA and the regulation of mRNA decay (Anderson and Parker, 1998;
Beilharz and Preiss, 2007; Gallie, 1991; Muhlrad et al., 1995; Munroe and Jacobson,
1990). All mRNAs undergo a deadenylation step regardless of whether or not the mRNA
is an NMD substrate (Norbury, 2013). The difference is that wild-type mRNA
degradation is initiated by deadenylation of the mRNA poly(A) tail, which is a
prerequisite for 5’ decapping of the mRNA (Decker and Parker, 1993; Muhlrad and
Parker, 1994). NMD substrates on the other hand undergo rapid 5’ decapping and rapid
deadenylation, but deadenylation is not required prior to decapping (Cao and Parker,
2003; Muhlrad et al., 1994; Muhlrad and Parker, 1994).
Deadenylation of an mRNA occurs through the combined efforts of the
178
Pan2/Pan3 and the Ccr4/Pop2/Not deadenylase complexes (Wahle and Winkler, 2013;
Wiederhold and Passmore, 2010). The Pan2 subunit of the Pan2/Pan3 complex exhibits
specific deadenylase activity and the Ccr4 subunit of the Ccr4/Pop2/Not is the catalytic
subunit (Chen et al., 2002; Daugeron et al., 2001; Tucker et al., 2002; Tucker et al.,
2001). There are conflicting reports on whether or not Caf1 is a catalytic subunit as well
in S. cerevisiae, although the catalytic activity of this subunit has been verified in other
organisms (Andersen et al., 2009; Daugeron et al., 2001; Jonstrup et al., 2007; Liang et
al., 2009; Thore et al., 2003; Tucker et al., 2002; Viswanathan et al., 2004). Previous
observations indicate that in the absence of the Pan2/Pan3 complex the Ccr4/Pop2/Not
complex is capable of full mRNA deadenylation (Tucker et al., 2001). Because
deadenylation is an important step in mRNA decay we hypothesized that deadenylation
would likely play a prominent role in the degradation of the YAP1 and GCN4 mRNAs.
To test this hypothesis we analyzed the YAP1 and GCN4 mRNA steady-state
accumulation in several deadenylation mutant strains. To our surprise, the various
deadenylation mutants did not cause a significant increase or decrease in the steady-state
accumulation of either the YAP1 or GCN4 mRNAs (Fig. 6). Because the Pan2/Pan3 and
Ccr4/Pop2/Not complexes can compensate for loss of the other it was not totally
surprising that single deletions which only effected one of the complexes did not result in
drastic changes in mRNA accumulation. However, deletions that render both complexes
inactive (ccr4Δpan2Δ) did not result in significant accumulation either (YAP1 mRNA
FCR ccr4Δpan2Δ/wild-type=1.1±0.1 and GCN4 mRNA FCR ccr4Δpan2Δ/wildtype=1.3±0.2; Fig. 6). The only strain in which there were slight notable differences,
179
although not considered significant according to the ≥ 2.0 FCR threshold, was the
ccr4Δcaf1Δ strain for the YAP1 mRNA (FCR ccr4Δcaf1Δ/wild-type=1.6±0.2) and in the
same strain for the GCN4 mRNA where the double deletion seemed to have the opposite
effect (FCR ccr4Δcaf1Δ/wild-type=0.6±0.0; Fig. 6). It is possible based on this
observation that the Pan2/Pan3 deadenylase complex is more efficient in compensating
for loss of Ccr4/Pop2/Not activity on the GCN4 mRNA than on the YAP1 mRNA.
We also analyzed steady-state accumulation of both the ENT4 mRNA and the PGK1
mRNA for comparison. Similar to the YAP1 and GCN4 mRNAs, neither the ENT4
mRNA nor the PGK1 mRNA seem to be largely effected by loss of deadenylation (Fig.
4). The only difference can be seen in the upf1Δ strains for the ENT4 mRNA, which is
expected since this mRNA is an NMD substrate.
From this we can conclude that blocking deadenylation through the Pan2/Pan3
complex or the Ccr4/Pop2/Not complex or both has little effect on YAP1 and GCN4
mRNA steady-state accumulation in cells. This can be explained by: 1) efficient
decapping and 5’3’ mRNA decay without prior deadenylation, and/or 2) another
mechanism of deadenylation aside from the Pan2/Pan3 and Ccr4/Pop2/Not deadenylase
complexes.
The YAP1 mRNA 3’ end is significantly shorter in xrn1Δ and dcp1Δ strains
compared to wild-type and upf1Δ strains
Finally, we wanted to determine at which step the YAP1 and GCN4 mRNAs are
deadenylated. We hypothesized that the mRNA would be likely deadenylated as both
180
wild-type mRNAs and NMD-substrates are deadenylated, but we were unsure if
deadenylation was occurring as a prerequisite to decapping. To do this we used the
Poly(A) Tail-Length Assay Kit from Affymetrix (Materials and Methods). Using this kit
a poly(A) polymerase first adds a short series of guanosine and inosine residues (G/I) to
the 3’ ends of polyadenylated mRNAs. The G/I tails become the priming site for reverse
transcription of the mRNAs to cDNA molecules. Two pairs of primers are then used to
amplify the 3’ ends of the desired mRNAs. The first primer pair is the gene-specific
(G.S.) primer pair, which consists of a forward and reverse primer that anneal to the 3’
end of the mRNA both of which are upstream of the polyadenylation site(s). It is
important to note that because the poly(A) sites for the mRNAs we used are not definitive
as there are multiple predicted poly(A) sites we designed the G.S. primer pair to anneal
upstream (3’) of all predicted poly(A) sites (SGD). The second primer pair is the poly(A)
tail (“Tail”) primer pair, which consists of the same upstream primer from the G.S.
primer pair and a universal reverse primer that anneals to the G/I tail (Fig. 7A). PCR
products obtained from the use of these two primer pairs are then compared in order to
determine the length of the 3’ poly(A) tail.
The Poly(A) Tail-Length Assay Kit was used to determine the poly(A) tail length
of steady-state YAP1, ENT4, and PGK1 mRNAs in wild-type, upf1Δ, xrn1Δ, and dcp1Δ
strains. Due to the cost of the kit and the number of reactions provided in each kit we did
not analyze the 3’ end of the GCN4 mRNA. An RNase H control was not done so we are
unable to positively confirm that the results observed are strictly due to differences in
poly(A) tail length. We can, however, use the results to analyze differences in the 3’ end
181
length of the mRNAs.
The results of this assay are shown in Figure 7B. In wild-type and upf1Δ strains
the YAP1 mRNA has a longer 3’ end compared to the xrn1Δ and dcp1Δ strains. This
pattern is consistent to what is seen for the PGK1 mRNA (not an NMD substrate) and
also the SSY5 mRNA (Chapter 3, Figure 5), which is a wild-type mRNA that is protected
from NMD (Fig. 7B; Kebaara and Atkin, 2009; Obenoskey et al., 2014; Chapter 2).
Additionally, the pattern of 3’ end length for the YAP1 mRNA is opposite of the pattern
for the ENT4 mRNA, which is a wild-type NMD substrate, in the tested strains.
However, it is important to note that the product resulting from use of the “Tail” primers
is smaller than the product for the G.S. primers, which should not be the case if the
primers were designed accurately. This indicates that the primers for the YAP1 mRNA
need redesigned for a more accurate analysis.
From this we can conclude that the YAP1 mRNA is able to be shortened at the 3’
end when either decapping or 5’3 decay are blocked. This also shows that the 3’ end
shortening is occurring much more slowly or not at all when both decapping and 5’3’
decay are functioning normally as in the wild-type and upf1Δ strains.
Discussion
Previously we presented the case of the SSY5 mRNA, which is a wild-type mRNA
in S. cerevisiae that contains multiple NMD-targeting signals but is not degraded by
NMD (Chapter 2). Here we investigate two additional wild-type mRNAs in S.
cerevisiae, the YAP1 and GCN4 mRNAs that also have at least one NMD-targeting
182
signal—a translated uORF—and are protected from degradation by NMD (Fig. 1). The
discovery of the YAP1 and GCN4 mRNAs as wild-type mRNAs protected from NMD
was not novel as this was previously shown (Ruiz-Echevarria et al., 1998; Vilela et al.,
1998). However, a follow-up study identified a simple mechanism of protection of these
two mRNAs from NMD by which trans-acting factor Pub1 binds to a stabilizer element
in the 5’ leader region of the mRNAs. In this study the binding of Pub1 to the STE was
convincingly shown to be solely responsible for the protection of both YAP1 and GCN4
mRNAs from NMD (Ruiz-Echevarria and Peltz, 2000). These results were of great
interest to us as we searched for the mechanism by which the SSY5 mRNA is protected
from degradation by NMD. The SSY5 mRNA also has a translated uORF just like the
YAP1 and GCN4 mRNAs, and it seemed like an appropriate hypothesis that Pub1 could
also be responsible for the protection of the SSY5 mRNA from NMD. However, when
we began the experiments to see if Pub1 was responsible for the protection of SSY5
mRNA from NMD (Chapter 2) we were unable to reproduce the results showing Pub1
was responsible for the protection of YAP1 and GCN4 mRNAs from NMD, which had
been included in the experiment as positive controls (Fig. 2). Cells were grown in both
rich and minimal media (Fig. 2) and we tested as many different strain backgrounds as
we could obtain in order to rule out all known variables (data not shown).
We were able to confirm that both the YAP1 and GCN4 mRNAs are likely
protected from degradation by NMD, but this stability is not quite as clear as previously
reported and is likely strain dependent (Figs. 1, 4-5). The T1/2 analysis of both mRNAs in
wild-type and upf1Δ strains is more indicative of protection from NMD than is the
183
analysis of the steady-state accumulation of the mRNAs in wild-type and upf1Δ strains
(Fig. 1). Further, we were able to show that YAP1 mRNA does cofractionate with heavy
fractions of the polyribosomes, which is an indication of association with polyribosomes
(Fig 3A). Likewise, the GCN4 mRNA is also found cofractionating where we would
expect although this pattern looks rather different from a normally translated mRNA (Fig.
3B). Translation of the GCN4 mRNA is regulated by the translation of its four short
uORFs, which are continually translated in order to repress the translation of the main
ORF until the gene product, Gcn4, is needed (Grant and Hinnebusch, 1994; Gunisova et
al., 2016; Hinnebusch, 2005). Thus, the GCN4 mRNA is expected to be present in the
lighter fractions since the continual translation of its four short uORFs means the mRNA
is associated with fewer ribosomes as these uORFs are only 2-3 codons in length.
Importantly, neither the YAP1 mRNA nor the GCN4 mRNA show differences in
fractionation between the wild-type and upf1Δ strains (Fig. 3A-B). Association with
polyribosomes (in the case of the YAP1 mRNA) or the few uORF occupying ribosomes
(in the case of the GCN4 mRNA) leads to the likely hypothesis that the mRNA is being
actively translated, although Westerns would need to be done in order to positively
confirm this. If the mRNA is being actively translated then it rules out the possibility that
the mRNA is protected from degradation by NMD by inhibition of translation.
We also looked at the stability of the YAP1 and GCN4 mRNAs in decapping,
5’3’ decay, and various deadenylation mutants. When 5’3’ decay is blocked in the
xrn1Δ strains, both the YAP1 and GCN4 mRNAs behave similarly and accumulate to
significantly higher levels compared to the wild-type strain. However, this accumulation
184
is not as great as what we saw for the SSY5 mRNA (Chapter 2) or for the ENT4 mRNA (a
wild-type NMD substrate; Fig. 4). The two most obvious explanations for this are 1) the
exosome-mediated 3’5’ decay of the YAP1 and GCN4 mRNAs is more efficient (i.e.
faster) than for the SSY5 or ENT4 mRNAs, or 2) Rat1 (also known as Kem1) digestion,
which was identified as the only other enzyme in yeast with 5’3’ exonuclease activity,
is able to work more efficiently on the YAP1 and GCN4 mRNAs than on the SSY5 or
ENT4 mRNAs (He and Jacobson, 2001).
A previous study also showed that Upf1 can actually stimulate the degradation of
decapped transcripts in an xrn1Δ strain (He and Jacobson, 2001). This could be one of
the reasons that we see higher steady-state accumulations and slightly longer T1/2 in the
xrn1Δupf1Δ strains. Another reason could also be that in this background the YAP1 and
GCN4 mRNAs are slightly more susceptible to NMD as can be seen comparing the wildtype and upf1Δ strains (Fig. 4).
Interestingly, blocking decapping in the dcp1Δ strain, which we expected to yield
a similar result as seen in the xrn1Δ strain, had very little influence on the steady-state
accumulation of either the YAP1 mRNA or the GCN4 mRNA (Fig. 5A). This indicates
that the exosome-mediated 3’5’ degradation of both mRNAs is much more efficient
when the 5’ mRNA cap is present since mRNAs in the xrn1Δ strain are decapped (He and
Jacobson, 2001). However, the T1/2 analysis for the YAP1 and GCN4 mRNAs in the
dcp1Δ strain tells a bit of a different story. Both mRNAs are more stable in the dcp1Δ
strain compared to the wild-type strain, and this increase in stability is significant for the
YAP1 mRNA (Fig. 5B). Taking into consideration the T1/2 analysis of both mRNAs in
185
the wild-type, xrn1Δ, and upf1Δ strains it does appear that blocking both decapping and
5’3’ degradation causes significant mRNA stabilization indicating that the 3’5’
degradation is not as efficient as the steady-state analysis in the dcp1Δ strain leads us to
believe. This also leads us to point out that steady-state accumulations do not always tell
the whole story and a T1/2 analysis is an important piece of the puzzle, which we also saw
in Figure 1.
When blocking deadenylation through deletion of the catalytic subunits of either
the Pan2/Pan3 complex or the Ccr4/Pop2/Not complex, or both we saw little influence on
steady-state mRNA accumulation of either the YAP1 mRNA or the GCN4 mRNA (Fig.
6). However, as pointed out previously, a T1/2 analysis of both mRNAs in these strains
would need to be done in order to make any definitive conclusions. If we make the
assumption that the T1/2 analysis would agree with the steady-state accumulations, then
we can say that blocking deadenylation has little effect on the degradation of the YAP1 or
GCN4 mRNAs. This tells us that deadenylation-independent decapping and subsequent
5’3’ decay is able to proceed in the absence of deadenylation. Although the Pan2/Pan3
complex and the Ccr4/Pop2/Not complex are able to compensate for loss of one another,
the fold changes in the double mutant ccr4Δpan2Δ are the most revealing about how lack
of deadenylation influences the stability of these two mRNAs (Fig. 6). To make this
analysis more complete it would also be helpful to look at the steady-state mRNA
accumulation and half-lives in 3’5’ decay (exosome) mutants.
When we used the Poly(A) Tail-Length Analysis kit to determine what was
happening on the 3’ end of the mRNA we found that the YAP1 mRNA behaved more
186
similarly to the PGK1 mRNA, which is not an NMD substrate and is typically degraded
5’3’ by the deadenylation-dependent decapping mechanism. However, the PGK1
mRNA can also undergo 3’5’ degradation when 5’3’ degradation is blocked, but this
process occurs more slowly (Muhlrad and Parker, 1994). This appears to be what is
happening for both the PGK1 mRNA and the YAP1 mRNA according to our results in
Figure 7. Both the PGK1 and YAP1 mRNAs show longer 3’ ends in the wild-type and
upf1Δ strains and shorter 3’ ends in the xrn1Δ and dcp1Δ strains. In contrast, the known
wild-type NMD substrate ENT4 mRNA shows the opposite pattern with longer 3’ ends in
the xrn1Δ and dcp1Δ strains than in the wild-type and upf1Δ strains (Fig. 7).
Going forward, it will be important to identify if the NMD mRNP, which
ultimately triggers the degradation of an mRNA through the NMD pathway, is
assembling on the YAP1 mRNA or the GCN4 mRNA. If the NMD mRNP is forming on
these mRNAs then protection from NMD is most likely incurred at a point downstream
of substrate recognition. Conversely, if the NMD mRNP complex never forms on the
mRNA, then protection is likely incurred before substrate recognition.
187
Figure Legends
Fig. 4-1. YAP1 mRNA is protected from degradation by NMD while GCN4 mRNA is
partially protected from degradation by NMD. A) Northern blot analysis of the steadystate accumulation of the YAP1, GCN4 and ENT4 (a wild-type NMD substrate) mRNAs
in wild-type (AAY277) and upf1Δ (AAY363) strains in the BY4741 background and in
wild-type (AAY187) and upf1Δ (AAY320) strains in the W303 background. Strains
were grown in rich media (YAPD). CYH2 pre-mRNA is an NMD control (CYH2 premRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown
for loading. Fold change values are normalized and determined as a fold change ratio
from wild-type; values are an average of three independent trials. B) Northern blot
analysis of the YAP1 and GCN4 mRNA half-lives in wild-type (AAY277) and upf1Δ
(AAY363) strains from the BY4741 background. Strains were grown in rich media
(YAPD). 10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Half-life
calculations were determined using a graph of percent mRNA remaining at each time
point during the exponential decay. C) Northern blot analysis of the steady-state
accumulation of the YAP1, GCN4 and ENT4 mRNAs in wild-type (AAY277) and upf1Δ
(AAY363) strains in the BY4741 background. Strains were grown in minimal media (SD
+ his, leu, met, ura, lys). D) Northern blot analysis of the YAP1 and GCN4 mRNA halflives in wild-type (AAY277) and upf1Δ (AAY363) strains from the BY4741 background.
Strains were grown in minimal media (SD + his, leu, met, ura, lys). 10µg/ml Thiolutin
was added to mid-log cultures at time=0 min. Half-life calculations were determined
using a graph of percent mRNA remaining at each time point during the exponential
188
decay and are the average of three independent trials.
Fig. 4-2. YAP1 and GCN4 mRNAs are not protected from NMD by trans-acting factor
Pub1. A) Northern blot analysis of the steady-state accumulation of YAP1 and GCN4
mRNAs in wild-type (AAY277), upf1Δ (AAY363), pub1Δ (AAY538), and pub1Δupf1Δ
(AAY590) strains in the BY4741 background. Strains were grown in rich media
(YAPD). CYH2 pre-mRNA is an NMD control (CYH2 pre-mRNA is an NMD substrate,
CYH2 mRNA is not an NMD substrate) and SCR1 is shown for loading. Fold change
values are normalized and determined as a fold change ratio from wild-type; values are
an average of three independent trials. B) Northern blot analysis of the steady-state
accumulation of YAP1 and GCN4 mRNAs in wild-type (AAY277), upf1Δ (AAY363),
and pub1Δ (AAY538) strains in the BY4741 background. Strains were grown in minimal
media (SD + his, leu, met, ura, lys). C) Northern blot analysis of YAP1 and GCN4
mRNA half-lives in wild-type (AAY277) and pub1Δ (AAY538) strains in the BY4741
background. Strains were grown in minimal media (SD + his, leu, met, ura, lys).
10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Half-life calculations
were determined using a graph of percent mRNA remaining at each time point during the
exponential decay and are the average of three independent trials.
Fig. 4-3. YAP1 and GCN4 mRNAs co-fractionate with polyribosomes. Polyribosome
analysis of YAP1 (A) and GCN4 (B) mRNAs in wild-type (AAY187) and upf1Δ
(AAY320) strains. Strains were grown in YAPD. Whole cell lysate was centrifuged
189
through a 15-50% sucrose gradient. Total RNA was extracted from each fraction
collected and total RNA from fractions 2-19 was transferred to a membrane for Northern
analysis. Fractions corresponding to the 40S ribosomal peak are in lanes 4-5, fractions
corresponding to the 60S ribosomal peak are in lanes 6-7, the fraction corresponding to
the 80S ribosomal peak is in lane 8 and polyribosome fractions are in lanes 9-15. The
graph represents the distribution of the mRNA in each fraction based on the relative
corrected volume of the mRNA present in each lane. In lanes where two bands were
present only the band corresponding to the size of the expected band for the mRNA was
quantified (~2.0 kb for the YAP1 mRNA). Polysome analysis was repeated twice with
similar results.
Fig. 4-4. YAP1 and GCN4 mRNAs accumulate significantly in xrn1Δ strains. Northern
blot analysis of the steady-state accumulation of the YAP1, GCN4, ENT4, and PGK1
mRNAs in wild-type (AAY360), upf1Δ (AAY589), xrn1Δ (AAY389), and xrn1Δupf1Δ
(AAY611) strains grown in YAPD. CYH2 pre-mRNA is an NMD control (CYH2 premRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown
for loading. Fold change values are normalized and determined as a fold change ratio
from wild-type; values are an average of three independent trials. B) mRNA half-life
analysis of the YAP1 and GCN4 mRNAs in the same strains used in part A. Strains were
grown in YAPD. 10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Halflife calculations were determined using a graph of percent mRNA remaining at each time
point during the exponential decay.
190
Fig. 4-5. YAP1 and GCN4 mRNAs do not accumulate in dcp1Δ strains, but half-life
analysis shows that dcp1Δ stabilizes the mRNAs albeit to different extents. A) Northern
blot analysis of the steady-state accumulation of the YAP1, GCN4, ENT4 and PGK1
mRNAs in wild-type (AAY360), upf1Δ (AAY589), dcp1Δ (AAY390), and dcp1Δupf1Δ
(AAY621) strains grown in YAPD. CYH2 pre-mRNA is an NMD control (CYH2 premRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown
for loading. Fold change values are normalized and determined as a fold change ratio
from wild-type; values are an average of three independent trials. B) mRNA half-life
analysis of the YAP1 and GCN4 mRNAs in the same strains used in part A. Strains were
grown in YAPD. 10µg/ml Thiolutin was added to mid-log cultures at time=0 min. Halflife calculations were determined using a graph of percent mRNA remaining at each time
point during the exponential decay.
Fig. 4-6. Deadenylation mutants show little effect on YAP1 and GCN4 mRNA stability.
Northern blot analysis of the steady-state accumulation of the YAP1, GCN4, ENT4, and
PGK1 mRNAs in wild-type (AAY391), upf1Δ (AAY594), ccr4Δ (AAY393), and
ccr4Δupf1Δ (AAY596), ccr4Δcaf1Δ (AAY394), ccr4Δcaf1Δupf1Δ (AAY609), pan2Δ
(AAY395), pan2Δupf1Δ (AAY595), pan2Δccr4Δ (AAY396), and pan2Δccr4Δupf1Δ
(AAY610) strains grown in YAPD. CYH2 pre-mRNA is an NMD control (CYH2 premRNA is an NMD substrate, CYH2 mRNA is not an NMD substrate) and SCR1 is shown
for loading. Fold change values are normalized and determined as a fold change ratio
191
from wild-type; values are an average of three independent trials.
Fig. 4-7. The YAP1 mRNA 3’ end is significantly shorter in xrn1Δ and dcp1Δ strains
compared to wild-type and upf1Δ strains. A) Schematic of how the Poly(A) Tail Length
Analysis works. This analysis was done using the “Poly(A) Tail-Length Assay Kit” from
Affymetrix (#76455). In this assay, a G/I tail is added to the end of mRNAs containing a
Poly(A) tail. These G/I-tailed mRNAs are then reverse transcribed to cDNA, which
provides a template for subsequent PCR. PCR is performed using two sets of primers
independently. The Gene Specific (G.S.; teal) primer pair is comprised of a forward
primer that binds anywhere within the open reading frame (ORF) or 3’ UTR and a
reverse primer that binds immediately upstream (5’) of the Poly(A) start site. *It is
important to note that the exact Poly(A) start site of the mRNAs in this figure were
unknown so the reverse primer was designed just upstream of the first predicted Poly(A)
start site according to the Saccharomyces Genome Database (SGD). The Poly(A) Tail
(Tail; pink) primer pair uses the same forward primer from the G.S. pair and a reverse
primer that anneals to the G/I tail (provided with the kit). B) Products from the PCR
reactions using the above primer pairs for the YAP1, ENT4, and PGK1 mRNAs in the
wild-type (AAY360), upf1Δ (AAY589), xrn1Δ (AAY389), and dcp1Δ (AAY390) strains
were resolved on a 2.5% agarose TAE gel. Strains were grown in YAPD and the total
RNA samples used for the reactions were the same RNA samples that were used for
Northern analysis in figures 2A and 3A.
192
Figure 4-1
A. Rich Media
B.
Rich Media
D. Minimal Media
C. Minimal Media
193
Figure 4-2
A. Rich Media
C. Minimal Media
B. Minimal Media
194
Figure 4-3
A.
B.
195
Figure 4-4
A.
B.
196
Figure 4-5
A.
B.
197
Figure 4-6
198
Figure 4-7
A.
B.
199
Table 4-1. Strains used in this study
Strain
Parent
Strain
Genotype
W303
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100
BY4741
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0
AAY320
W303
AAY360
AAY363
BY4741
AAY389
AAY360
AAY390
AAY360
AAY391
AAY393
AAY391
AAY394
AAY391
AAY395
AAY391
AAY396
AAY391
AAY538
BY4741
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100 upf1::URA3
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 upf1::URA3
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100 xrn1::ADE2
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100 dcp1::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
cup1Δ::LEU2PM
MATa his4-539 trp1 ura3-52 leu2-3,112
cup1Δ::LEU2PM ccr4::NEO
MATa his4-539 trp1 ura3-52 leu2-3,112
cup1Δ::LEU2PM ccr4::NEO caf1::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
cup1Δ::LEU2PM pan2::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
cup1Δ::LEU2PM ccr4::NEO pan2::URA3
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 pub1Δ
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1100 upf1::URA3
MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 pub1Δ
AAY590* AAY538
upf1::URA3
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY594* AAY391
cup1Δ::LEU2PM upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY595* AAY395
cup1Δ::LEU2PM pan2::URA3 upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY596* AAY393
cup1Δ::LEU2PM ccr4::NEO upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY609* AAY394
cup1Δ::LEU2PM ccr4::NEO caf1::URA3 upf1::TRP1
MATa his4-539 trp1 ura3-52 leu2-3,112
AAY610* AAY396
cup1Δ::LEU2PM ccr4::NEO pan2::URA3 upf1::TRP1
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY611* AAY389
100 xrn1::ADE2 upf1::TRP1
MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1AAY621* AAY390
100 dcp1::URA3 upf1::TRP1
* = new strain created for this study
AAY589*
AAY360
Source
Ralser et al.,
2012
Winston et al.,
1998
Atkin lab
He and
Jacobson, 1995
Atkin lab
He and
Jacobson, 2001
He and
Jacobson, 2001
Tucker et al.,
2001
Tucker et al.,
2001
Tucker et al.,
2001
Tucker et al.,
2001
Tucker et al.,
2001
Open
Biosystems
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
Atkin lab
200
References
Amrani, N., Dong, S., He, F., Ganesan, R., Ghosh, S., Kervestin, S., Li, C., Mangus,
D.A., Spatrick, P., and Jacobson, A. (2006). Aberrant termination triggers nonsensemediated mRNA decay. Biochem Soc Trans 34, 39-42.
Amrani, N., Ganesan, R., Kervestin, S., Mangus, D.A., Ghosh, S., and Jacobson, A.
(2004). A faux 3'-UTR promotes aberrant termination and triggers nonsense-mediated
mRNA decay. Nature 432, 112-118.
Andersen, K.R., Jonstrup, A.T., Van, L.B., and Brodersen, D.E. (2009). The activity and
selectivity of fission yeast Pop2p are affected by a high affinity for Zn2+ and Mn2+ in
the active site. RNA 15, 850-861.
Anderson, J.S., and Parker, R.P. (1998). The 3' to 5' degradation of yeast mRNAs is a
general mechanism for mRNA turnover that requires the SKI2 DEVH box protein and 3'
to 5' exonucleases of the exosome complex. EMBO J 17, 1497-1506.
Atkin, A.L., Altamura, N., Leeds, P., and Culbertson, M.R. (1995). The majority of yeast
UPF1 co-localizes with polyribosomes in the cytoplasm. Mol Biol Cell 6, 611-625.
Barbosa, C., Peixeiro, I., and Romao, L. (2013). Gene expression regulation by upstream
open reading frames and human disease. PLoS Genet 9, e1003529.
Bedwell, D.M., Kaenjak, A., Benos, D.J., Bebok, Z., Bubien, J.K., Hong, J., Tousson, A.,
Clancy, J.P., and Sorscher, E.J. (1997). Suppression of a CFTR premature stop mutation
in a bronchial epithelial cell line. Nat Med 3, 1280-1284.
Beelman, C.A., Stevens, A., Caponigro, G., LaGrandeur, T.E., Hatfield, L., Fortner,
D.M., and Parker, R. (1996). An essential component of the decapping enzyme required
for normal rates of mRNA turnover. Nature 382, 642-646.
Behm-Ansmant, I., Gatfield, D., Rehwinkel, J., Hilgers, V., and Izaurralde, E. (2007). A
conserved role for cytoplasmic poly(A)-binding protein 1 (PABPC1) in nonsensemediated mRNA decay. EMBO J 26, 1591-1601.
Beilharz, T.H., and Preiss, T. (2007). Widespread use of poly(A) tail length control to
accentuate expression of the yeast transcriptome. RNA 13, 982-997.
Brown, C.E., and Sachs, A.B. (1998). Poly(A) tail length control in Saccharomyces
cerevisiae occurs by message-specific deadenylation. Mol Cell Biol 18, 6548-6559.
Cao, D., and Parker, R. (2003). Computational modeling and experimental analysis of
nonsense-mediated decay in yeast. Cell 113, 533-545.
Chen, J., Chiang, Y.C., and Denis, C.L. (2002). CCR4, a 3'-5' poly(A) RNA and ssDNA
exonuclease, is the catalytic component of the cytoplasmic deadenylase. EMBO J 21,
1414-1426.
201
Coller, J., and Parker, R. (2004). Eukaryotic mRNA decapping. Annu Rev Biochem 73,
861-890.
Conti, E., and Izaurralde, E. (2005). Nonsense-mediated mRNA decay: molecular
insights and mechanistic variations across species. Curr Opin Cell Biol 17, 316-325.
Cui, Y., Hagan, K.W., Zhang, S., and Peltz, S.W. (1995). Identification and
characterization of genes that are required for the accelerated degradation of mRNAs
containing a premature translational termination codon. Genes Dev 9, 423-436.
Daugeron, M.C., Mauxion, F., and Seraphin, B. (2001). The yeast POP2 gene encodes a
nuclease involved in mRNA deadenylation. Nucleic Acids Res 29, 2448-2455.
Decker, C.J., and Parker, R. (1993). A turnover pathway for both stable and unstable
mRNAs in yeast: evidence for a requirement for deadenylation. Genes Dev 7, 1632-1643.
Gallie, D.R. (1991). The cap and poly(A) tail function synergistically to regulate mRNA
translational efficiency. Genes Dev 5, 2108-2116.
Gietz, R.D., and Woods, R.A. (2002). Transformation of yeast by lithium acetate/singlestranded carrier DNA/polyethylene glycol method. Methods Enzymol 350, 87-96.
Gonzalez, C.I., Bhattacharya, A., Wang, W., and Peltz, S.W. (2001). Nonsense-mediated
mRNA decay in Saccharomyces cerevisiae. Gene 274, 15-25.
Grant, C.M., and Hinnebusch, A.G. (1994). Effect of sequence context at stop codons on
efficiency of reinitiation in GCN4 translational control. Mol Cell Biol 14, 606-618.
Guan, Q., Zheng, W., Tang, S., Liu, X., Zinkel, R.A., Tsui, K.W., Yandell, B.S., and
Culbertson, M.R. (2006). Impact of nonsense-mediated mRNA decay on the global
expression profile of budding yeast. PLoS Genet 2, e203.
Gunisova, S., Beznoskova, P., Mohammad, M.P., Vlckova, V., and Valasek, L.S. (2016).
In-depth analysis of cis-determinants that either promote or inhibit reinitiation on GCN4
mRNA after translation of its four short uORFs. RNA 22, 542-558.
Hagan, K.W., Ruiz-Echevarria, M.J., Quan, Y., and Peltz, S.W. (1995). Characterization
of cis-acting sequences and decay intermediates involved in nonsense-mediated mRNA
turnover. Mol Cell Biol 15, 809-823.
Hall, G.W., and Thein, S. (1994). Nonsense codon mutations in the terminal exon of the
beta-globin gene are not associated with a reduction in beta-mRNA accumulation: a
mechanism for the phenotype of dominant beta-thalassemia. Blood 83, 2031-2037.
He, F., Brown, A.H., and Jacobson, A. (1997). Upf1p, Nmd2p, and Upf3p are interacting
components of the yeast nonsense-mediated mRNA decay pathway. Mol Cell Biol 17,
1580-1594.
202
He, F., and Jacobson, A. (1995). Identification of a novel component of the nonsensemediated mRNA decay pathway by use of an interacting protein screen. Genes Dev 9,
437-454.
He, F., and Jacobson, A. (2001). Upf1p, Nmd2p, and Upf3p regulate the decapping and
exonucleolytic degradation of both nonsense-containing mRNAs and wild-type mRNAs.
Mol Cell Biol 21, 1515-1530.
He, F., Li, X., Spatrick, P., Casillo, R., Dong, S., and Jacobson, A. (2003). Genome-wide
analysis of mRNAs regulated by the nonsense-mediated and 5' to 3' mRNA decay
pathways in yeast. Mol Cell 12, 1439-1452.
He, F., Peltz, S.W., Donahue, J.L., Rosbash, M., and Jacobson, A. (1993). Stabilization
and ribosome association of unspliced pre-mRNAs in a yeast upf1- mutant. Proc Natl
Acad Sci U S A 90, 7034-7038.
Hentze, M.W., and Kulozik, A.E. (1999). A perfect message: RNA surveillance and
nonsense-mediated decay. Cell 96, 307-310.
Hilleren, P., McCarthy, T., Rosbash, M., Parker, R., and Jensen, T.H. (2001). Quality
control of mRNA 3'-end processing is linked to the nuclear exosome. Nature 413, 538542.
Hinnebusch, A.G. (2005). Translational regulation of GCN4 and the general amino acid
control of yeast. Annu Rev Microbiol 59, 407-450.
Hsu, C.L., and Stevens, A. (1993). Yeast cells lacking 5'-->3' exoribonuclease 1 contain
mRNA species that are poly(A) deficient and partially lack the 5' cap structure. Mol Cell
Biol 13, 4826-4835.
Isken, O., and Maquat, L.E. (2007). Quality control of eukaryotic mRNA: safeguarding
cells from abnormal mRNA function. Genes Dev 21, 1833-1856.
Isken, O., and Maquat, L.E. (2008). The multiple lives of NMD factors: balancing roles
in gene and genome regulation. Nat Rev Genet 9, 699-712.
Johansson, M.J., He, F., Spatrick, P., Li, C., and Jacobson, A. (2007). Association of
yeast Upf1p with direct substrates of the NMD pathway. Proc Natl Acad Sci U S A 104,
20872-20877.
Jonstrup, A.T., Andersen, K.R., Van, L.B., and Brodersen, D.E. (2007). The 1.4-A crystal
structure of the S. pombe Pop2p deadenylase subunit unveils the configuration of an
active enzyme. Nucleic Acids Res 35, 3153-3164.
Kebaara, B., Nazarenus, T., Taylor, R., and Atkin, A.L. (2003). Genetic background
affects relative nonsense mRNA accumulation in wild-type and upf mutant yeast strains.
Curr Genet 43, 171-177.
203
Kebaara, B.W., and Atkin, A.L. (2009). Long 3'-UTRs target wild-type mRNAs for
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Nucleic Acids Res 37,
2771-2778.
Kebaara, B.W., Baker, K.E., Patefield, K.D., and Atkin, A.L. (2012). Analysis of
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Curr Protoc Cell Biol
Chapter 27, Unit 27 23.
Lee, B.S., and Culbertson, M.R. (1995). Identification of an additional gene required for
eukaryotic nonsense mRNA turnover. Proc Natl Acad Sci U S A 92, 10354-10358.
Lelivelt, M.J., and Culbertson, M.R. (1999). Yeast Upf proteins required for RNA
surveillance affect global expression of the yeast transcriptome. Mol Cell Biol 19, 67106719.
Lewis, B.P., Green, R.E., and Brenner, S.E. (2003). Evidence for the widespread
coupling of alternative splicing and nonsense-mediated mRNA decay in humans. Proc
Natl Acad Sci U S A 100, 189-192.
Liang, W., Li, C., Liu, F., Jiang, H., Li, S., Sun, J., Wu, X., and Li, C. (2009). The
Arabidopsis homologs of CCR4-associated factor 1 show mRNA deadenylation activity
and play a role in plant defence responses. Cell Res 19, 307-316.
Lykke-Andersen, J., Shu, M.D., and Steitz, J.A. (2001). Communication of the position
of exon-exon junctions to the mRNA surveillance machinery by the protein RNPS1.
Science 293, 1836-1839.
Maderazo, A.B., He, F., Mangus, D.A., and Jacobson, A. (2000). Upf1p control of
nonsense mRNA translation is regulated by Nmd2p and Upf3p. Mol Cell Biol 20, 45914603.
Mangus, D.A., Evans, M.C., and Jacobson, A. (2003). Poly(A)-binding proteins:
multifunctional scaffolds for the post-transcriptional control of gene expression. Genome
Biol 4, 223.
Mangus, D.A., and van Hoof, A. (2003). Making and breaking the message. Genome
Biol 4, 346.
Maquat, L.E., and Carmichael, G.G. (2001). Quality control of mRNA function. Cell 104,
173-176.
Maquat, L.E., and Serin, G. (2001). Nonsense-mediated mRNA decay: insights into
mechanism from the cellular abundance of human Upf1, Upf2, Upf3, and Upf3X
proteins. Cold Spring Harb Symp Quant Biol 66, 313-320.
McGlincy, N.J., and Smith, C.W. (2008). Alternative splicing resulting in nonsensemediated mRNA decay: what is the meaning of nonsense? Trends Biochem Sci 33, 385393.
204
Mendell, J.T., Sharifi, N.A., Meyers, J.L., Martinez-Murillo, F., and Dietz, H.C. (2004).
Nonsense surveillance regulates expression of diverse classes of mammalian transcripts
and mutes genomic noise. Nat Genet 36, 1073-1078.
Mitchell, P., and Tollervey, D. (2003). An NMD pathway in yeast involving accelerated
deadenylation and exosome-mediated 3'-->5' degradation. Mol Cell 11, 1405-1413.
Muhlemann, O., Eberle, A.B., Stalder, L., and Zamudio Orozco, R. (2008). Recognition
and elimination of nonsense mRNA. Biochim Biophys Acta 1779, 538-549.
Muhlrad, D., Decker, C.J., and Parker, R. (1994). Deadenylation of the unstable mRNA
encoded by the yeast MFA2 gene leads to decapping followed by 5'-->3' digestion of the
transcript. Genes Dev 8, 855-866.
Muhlrad, D., Decker, C.J., and Parker, R. (1995). Turnover mechanisms of the stable
yeast PGK1 mRNA. Mol Cell Biol 15, 2145-2156.
Muhlrad, D., and Parker, R. (1994). Premature translational termination triggers mRNA
decapping. Nature 370, 578-581.
Muhlrad, D., and Parker, R. (1999). Aberrant mRNAs with extended 3' UTRs are
substrates for rapid degradation by mRNA surveillance. RNA 5, 1299-1307.
Munroe, D., and Jacobson, A. (1990). Tales of poly(A): a review. Gene 91, 151-158.
Ni, J.Z., Grate, L., Donohue, J.P., Preston, C., Nobida, N., O'Brien, G., Shiue, L., Clark,
T.A., Blume, J.E., and Ares, M., Jr. (2007). Ultraconserved elements are associated with
homeostatic control of splicing regulators by alternative splicing and nonsense-mediated
decay. Genes Dev 21, 708-718.
Norbury, C.J. (2013). Cytoplasmic RNA: a case of the tail wagging the dog. Nat Rev Mol
Cell Biol 14, 643-653.
Nyiko, T., Sonkoly, B., Merai, Z., Benkovics, A.H., and Silhavy, D. (2009). Plant
upstream ORFs can trigger nonsense-mediated mRNA decay in a size-dependent manner.
Plant Mol Biol 71, 367-378.
Obenoskey, J., Lane, D.R., Atkin, A.L., and Kebaara, B.W. (2014). Immunity of the
Saccharomyces cerevisiae SSY5 mRNA to nonsense-mediated mRNA decay. Front Mol
Biosci 1, 25.
Oliveira, C.C., and McCarthy, J.E. (1995). The relationship between eukaryotic
translation and mRNA stability. A short upstream open reading frame strongly inhibits
translational initiation and greatly accelerates mRNA degradation in the yeast
Saccharomyces cerevisiae. J Biol Chem 270, 8936-8943.
Plant, E.P., Wang, P., Jacobs, J.L., and Dinman, J.D. (2004). A programmed -1 ribosomal
frameshift signal can function as a cis-acting mRNA destabilizing element. Nucleic Acids
Res 32, 784-790.
205
Pulak, R., and Anderson, P. (1993). mRNA surveillance by the Caenorhabditis elegans
smg genes. Genes Dev 7, 1885-1897.
Ruiz-Echevarria, M.J., Gonzalez, C.I., and Peltz, S.W. (1998). Identifying the right stop:
determining how the surveillance complex recognizes and degrades an aberrant mRNA.
EMBO J 17, 575-589.
Ruiz-Echevarria, M.J., and Peltz, S.W. (2000). The RNA binding protein Pub1 modulates
the stability of transcripts containing upstream open reading frames. Cell 101, 741-751.
Shoemaker, C.J., and Green, R. (2012). Translation drives mRNA quality control. Nat
Struct Mol Biol 19, 594-601.
Thore, S., Mauxion, F., Seraphin, B., and Suck, D. (2003). X-ray structure and activity of
the yeast Pop2 protein: a nuclease subunit of the mRNA deadenylase complex. EMBO
Rep 4, 1150-1155.
Tucker, M., Staples, R.R., Valencia-Sanchez, M.A., Muhlrad, D., and Parker, R. (2002).
Ccr4p is the catalytic subunit of a Ccr4p/Pop2p/Notp mRNA deadenylase complex in
Saccharomyces cerevisiae. EMBO J 21, 1427-1436.
Tucker, M., Valencia-Sanchez, M.A., Staples, R.R., Chen, J., Denis, C.L., and Parker, R.
(2001). The transcription factor associated Ccr4 and Caf1 proteins are components of the
major cytoplasmic mRNA deadenylase in Saccharomyces cerevisiae. Cell 104, 377-386.
Vilela, C., Linz, B., Rodrigues-Pousada, C., and McCarthy, J.E. (1998). The yeast
transcription factor genes YAP1 and YAP2 are subject to differential control at the levels
of both translation and mRNA stability. Nucleic Acids Res 26, 1150-1159.
Viswanathan, P., Ohn, T., Chiang, Y.C., Chen, J., and Denis, C.L. (2004). Mouse CAF1
can function as a processive deadenylase/3'-5'-exonuclease in vitro but in yeast the
deadenylase function of CAF1 is not required for mRNA poly(A) removal. J Biol Chem
279, 23988-23995.
Wahle, E., and Winkler, G.S. (2013). RNA decay machines: deadenylation by the Ccr4not and Pan2-Pan3 complexes. Biochim Biophys Acta 1829, 561-570.
Welch, E.M., and Jacobson, A. (1999). An internal open reading frame triggers nonsensemediated decay of the yeast SPT10 mRNA. EMBO J 18, 6134-6145.
Wiederhold, K., and Passmore, L.A. (2010). Cytoplasmic deadenylation: regulation of
mRNA fate. Biochem Soc Trans 38, 1531-1536.
206
CHAPTER 5
N5-Phosphonoacetyl-L-ornithine (PALO): influence on regulation of amino acid
biosynthetic genes in Saccharomyces cerevisiae
Reference:
Johnson, B., Steadman, R., Patefield, K.D., Bunker, J.J., Atkin, A.L., Dussalt, P.
(2011) N5-Phosphonoacetyl-L-ornithine (PALO): A convenient synthesis and
investigation of influence on regulation of amino acid biosynthetic genes in
Saccharomyces cerevisiae. Bioinorganic and Med. Chem. Letters. 21: 2351-2353.
207
Abstract
A scalable four-step synthesis of the ornithine transcarbamylase inhibitor N5phosphonoacetyl-L-ornithine (PALO) is reported based upon selective protection of the
amino acid of ornithine as a boroxazolidinone. Investigations in the model organism
Saccharomyces cerevisiae found PALO did not influence growth rate or expression of
genes involved in arginine metabolism.
208
Introduction
N5-Phosphonoacetyl-L-ornithine (PALO) is a bisubstrate transition-state analog
which competitively inhibits ornithine transcarbamylase (OTC), shutting down the
biosynthesis of citrulline in both the urea and the arginine biosynthetic pathways (Mori et
al., 1977; Penninckx and Gigot, 1978). Competitive inhibitors of amino acid metabolism
have been key components in studies ranging from arginine starvation in fungi and bacteria
to studies on OTC deficiency (Alewood et al., 1984; Kinney and Lusty, 1989).
Earlier work by Kinney and Lusty suggested that PALO would be an effective tool
for the study of arginine metabolism in the yeast Saccharomyces cerevisiae (Kinney and
Lusty, 1989). Arginine is used by all organisms for protein synthesis, and is also used by
yeast as a nitrogen source (Hoogenraad, 1978; Penninckx and Gigot, 1978, 1979). The
biosynthesis and metabolism of arginine are carefully regulated (Wu and Morris, 1998).
Arginine metabolism is required for the escape of the human pathogenic yeast Candida
albicans from macrophages (Lorenz et al., 2004). We were specifically interested in
applying PALO as a tool to determine whether both arginine biosynthesis and metabolism
are required for this evasion of the immune system, which is presumed critical to the
success of C. albicans as an invasive opportunistic pathogen. We now report a short and
scalable synthesis of PALO, as well as results of investigations into the influence of PALO
on growth and arginine metabolism in the model organism S. cerevisiae.
Materials and Methods
Yeast strains and growth conditions
209
Saccharomyces cerevisiae strains BY4741 (MATa his3-1 leu2-0 met15-0 ura3-0)
(Ref 1, 2, 7), YSC1178-7500224 (MATa his3-1 leu2-0 met15-0 ura3-0 ARG5,6TAP), YSC1178-7502950 (MATa his3-1 leu2-0 met15-0 ura3-0, CPA1-TAP) and
YSC1178-7500415 (MATa his3-1 leu2-0 met15-0 ura3-0, TRP5-TAP) were used.
BY4741 was purchased from the American Type Culture Collection (ATCC, Manassas,
VA). The YSC1178 strains were purchased from Open Biosystems Products (Huntsville,
AL). S. cerevisiae strains were inoculated from a saturated culture incubated at 30°C in
YPD in a tube roller in minimal medium supplemented with leucine, methionine, uracil
and histidine. Cultures were treated with 15 µM PALO in a 50 mM potassium phosphate
pH 6.8 buffer, potassium phosphate buffer pH 6.8 alone (untreated), 1 mg/mL arginine,
or both 15 µM PALO and 1 mg/mL arginine at the time of inoculation. Cells were
incubated at 30°C and then harvested at mid-log growth phase (OD600 0.4-0.6). Cells
were observed for effects on growth via incubation at 30°C in nitrogen-limiting medium
containing 2% dextrose, 1 mM ammonium sulfate, 100 µM potassium phosphate buffer
pH 6.8, and 10 µg/mL leucine, methionine, uracil and histidine.
Quantitative northern analysis of mRNA levels
Steady-state mRNA levels were measured as previously described. Oligolabeled DNA
probes were used to probe the northern blots. DNA probes were generated using primer
sets for amplifying yeast open reading frames based on the sequences available from the
Saccharomyces Genome Database (http://www.yeastgenome.org/).
210
Western blot analysis
Western blots were prepared as previously described (Atkin et al., 1995). TAP-tagged
Arg5,6p, Cpa1p and Trp5p were detected with Supersignal West Pico chemiluminescent
substrate, using the manufacturers protocol (Pierce, Rockford, IL). Rabbit anti-TAP
polyclonal antibodies were purchased from Thermo Fisher Scientific Inc. (Rockford, IL)
and goat anti-rabbit IgG polyclonal antibodies conjugated with horseradish peroxidase
were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).
Results
A study by Kinney and Lusty reported that PALO increased expression of CPA1,
CPA2, HIS3, and TRP5 reporter constructs in the yeast S. cerevisiae (Kinney and Lusty,
1989). The data were consistent with limitation of arginine leading to specific up
regulation of arginine biosynthetic genes (CPA1 and CPA2), and a general amino acid
starvation response typified by up regulation of HIS3 and TRP5.
We tested whether treatment with PALO resulted in changes in mRNA
accumulation and/or translation of genes involved in amino acid biosynthesis. We
examined expression of two genes required for arginine biosynthesis; ARG5,6, which
encodes protein involved in the second and third steps of arginine biosynthesis from
glutamate, and CPA1, which encodes a subunit of arginine-specific carbamyl-phosphate
synthetase. Both of these genes are up regulated specifically in response to arginine
limitation. Expression of TRP5, which encodes an enzyme that catalyzes the final step in
tryptophan biosynthesis, was also examined as an indicator of the general amino acid
211
starvation response. Quantitative northern analysis was performed to measure mRNA
abundance, a measure of the combined rates of transcription and mRNA decay. Western
analysis was performed to measure mRNA abundance, a measure of the combined rates
of transcription and MRNA decay. Western Analysis was used to look at the level of
protein synthesis for these genes. This combined approach enabled direct evaluation of
effects on transcription, mRNA stability, and translation.
Although the yeast strains from the earlier studies were not available, (Kinney and
Lusty, 1989) they were derived from the W303 genetic background, which usually carry
the can1-100 mutation (Thomas and Rothstein, 1989). As CAN1 encodes an arginine
permease required for efficient uptake of arginine, the can1-100 mutation is not well
suited for the current experiments. As a compromise, and to assess if PALO will be
generally useful as a metabolic tool in S. cerevisiae, the current data was obtained using
the BY4741 strain of S. cerevisiae (Brachmann et al., 1998). This strain is derived from
S288C, the strain used in the systematic sequencing project, and it is the most commonly
used genetic background for genome-wide functional analyses.
If PALO affects transcription or mRNA stability, we expected to observe an
increase in expression of ARG5,6 CPA1, and TRP5 mRNA transcripts in the presence of
PALO. We also expected that addition of arginine would relieve the arginine limitation
induced by PALO. Cells were grown and treated with either 15 μM of the synthesized
PALO, 1 mg/mL L-arginine, or 15 μM of the synthesized PALO and 1 mg/mL L-arginine
as described by Kinney and Lusty (Kinney and Lusty, 1989). An untreated culture was
grown in the same media as a control. The abundance of ARG5,6, CPA1, and TRP5
212
mRNAs was measured by quantitative northern analysis (Fig. 1A). The abundance of
ARG5,6, CPA1, and TRP5 mRNAs in the PALO treated cells was not significantly
different from the untreated cells. The abundance of ARG5,6 and CPA1, but not TRP5
mRNAs was lower in cells treated with arginine regardless of whether the cells were
treated with PALO or not. Additionally, the abundance of the TRP5 transcript did not
show any variation in response to addition of PALO, arginine, or both. In summary,
treatment with PALO had no effect on the abundance of ARG5,6, CPA1 or TRP5
mRNAs.
The possible effect of PALO on translation was investigated by western analysis
(Fig. 1B). If PALO affects translation, we expected to see an increase in the production
of Arg5,6p, Cpa1p, and Trp5p in the presence of PALO. We also expected that addition
of L-arginine would reduce protein levels of Arg5,6p and Cpa1p, but not Trp5p. We
used cell extracts from strains (YSC1178-7500224, YSC1178-7502950 and YSC11787500415, respectively) carrying TAP-tagged alleles of ARG5,6, CPA1, and TRP5 for the
Western analysis. These strains produce a TAP-fusion protein for each gene, which can
be detected on western blots with anti-TAP antibodies. BY4741 was used as an untagged
control because it is the parent strain for the TAP-tagged strains. Cells were grown and
treated in the same manner as for the northern analysis. Measurement of the signal
intensity of the bands was detected by western blotting revealed no difference in the
production of Arg5,6, Cpa1, or Trp5 between strains treated with 15 μM PALO and the
control. As expected, addition of L-arginine reduced production of Arg5,6 and Cpa1p,
but not Trp5p regardless of whether PALO was added or not. In summary, PALO had no
213
effect on Arg5,6, Cpa1, and Trp5 protein levels.
PALO did not affect the growth rate of BY4741 in minimal media or nitrogenlimiting medium (data not shown), used in an attempt to increase sensitivity to PALO by
lowering intracellular arginine concentrations. Growth in nitrogen-limiting medium
induces cells to use amino acids, including arginine, as a source of nitrogen. These
findings were consistent with the observations reported by Kinney and Lusty (Kinney and
Lusty, 1989).
Conclusions
We have developed an efficient synthesis of very pure PALO by a route suitable
for both analytical and preparative applications. In contrast to a previous report
describing PALO-induced arginine starvation, (Kinney and Lusty, 1989) we observed no
response in S. cerevisiae strain BY4741 to treatment with PALO (Fig. 1). The
discrepancy suggests either that not all yeast strains are sensitive to PALO or that the
earlier indications of arginine starvation resulted from impurities in the samples of PALO
employed. The current results are consistent with observations that PALO had no effect
in isolated rat mitochondria, intact rat hepatocytes, or E. coli (Hoogenraad, 1978;
Penninckx and Gigot, 1979).
214
Figure Legend
Fig. 5-1. PALO has no effect on expression of the ARG5,6, CPA1, and TRP5 genes in
Saccharomyces cerevisiae. S. cerevisiae strains (Brachmann et al., 1998) were
inoculated from a saturated culture into minimal medium supplemented with leucine,
methionine, uracil, and histidine. Cultures were treated with 15 μM PALO in a potassium
phosphate pH 6.8 buffer, potassium phosphate buffer pH 6.8 alone (untreated), 1 mg/mL
L-Arginine, or both 15 μM PALO and 1 mg/mL L-Arginine at the time of inoculation.
Cells were incubated at 30°C and then harvested at mid-log growth phase (OD600 0.40.6). A) Northern blots were prepared with total RNA extracted from BY4741
(Brachmann et al., 1998) and probed with oligolabeled DNA probes (Kebaara et al.,
2003). DNA probes were generated using primer sets for amplifying yeast open reading
frames based on the sequences available from the Saccharomyces Genome Database.
Shown are representative phosphor-images of a northern blot probed with radioactive
ARG5,6, CPA1, TRP5, and SCR1 DNAs. SCR1 was used as a loading control.
Quantitative results depicted are the average of three replicate trials and are normalized to
the SCR1 loading control. B) Western blots were prepared using protein extracts from S.
cerevisiae YSC1178-7500224, YSC1178-7502950 and YSC1178-7500415 strains
expressing TAP-fusion proteins Arg5,6-TAP, Cpa1-TAP, and Trp5-TAP, respectively, as
well as BY4741 (untagged control; Atkin et al., 1995). The TAP-tagged proteins were
detected using an anti-TAP antibody. No proteins bound the anti-TAP antibody in the
untagged control. Duplicate polyacrylamide gels were stained with Coomassie blue for
use as loading controls. The corresponding loading controls are shown beneath the
215
western blots.
216
Figure 5-1
217
References
Alewood, P.F., Johns, R.B., Hoogenraad, N.J., and Sutherland, T. (1984). An Improved
Synthesis of Nδ-(Phosphonoacetyl)-L-ornithine. Synthesis 1984, 403-404.
Atkin, A.L., Altamura, N., Leeds, P., and Culbertson, M.R. (1995). The majority of yeast
UPF1 co-localizes with polyribosomes in the cytoplasm. Mol Biol Cell 6, 611-625.
Brachmann, C.B., Davies, A., Cost, G.J., Caputo, E., Li, J., Hieter, P., and Boeke, J.D.
(1998). Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful
set of strains and plasmids for PCR-mediated gene disruption and other applications.
Yeast 14, 115-132.
Hoogenraad, N.J. (1978). Synthesis and properties of δ-N-(phosphonacetyl)-l-ornithine.
Archives of Biochemistry and Biophysics 188, 137-144.
Kebaara, B., Nazarenus, T., Taylor, R., and Atkin, A.L. (2003). Genetic background
affects relative nonsense mRNA accumulation in wild-type and upf mutant yeast strains.
Curr Genet 43, 171-177.
Kinney, D.M., and Lusty, C.J. (1989). Arginine restriction induced by delta-N(phosphonacetyl)-L-ornithine signals increased expression of HIS3, TRP5, CPA1, and
CPA2 in Saccharomyces cerevisiae. Mol Cell Biol 9, 4882-4888.
Lorenz, M.C., Bender, J.A., and Fink, G.R. (2004). Transcriptional response of Candida
albicans upon internalization by macrophages. Eukaryot Cell 3, 1076-1087.
Mori, M., Aoyagi, K., Tatibana, M., Ishikawa, T., and Ishii, H. (1977). N delta(phosphonacetyl)-L-ornithine, a potent transition state analogue inhibitor of ornithine
carbamoyltransferase. Biochem Biophys Res Commun 76, 900-904.
Penninckx, M., and Gigot, D. (1978). Synthesis and interaction with Escherichia coli Lornithine carbamolytransferase of two potential transition-state analogues. FEBS Lett 88,
94-96.
Penninckx, M., and Gigot, D. (1979). Synthesis of a peptide form of N-delta(phosphonoacetyl)-L-ornithine. Its antibacterial effect through the specific inhibition of
Escherichia coli L-ornithine carbamoyltransferase. J Biol Chem 254, 6392-6396.
Thomas, B.J., and Rothstein, R. (1989). Elevated recombination rates in transcriptionally
active DNA. Cell 56, 619-630.
Wu, G., and Morris, S.M., Jr. (1998). Arginine metabolism: nitric oxide and beyond.
Biochem J 336 ( Pt 1), 1-17.
218
CHAPTER 6:
Summary and Future Directions
219
Summary and Future Directions
The regulation of gene expression in eukaryotes is a very delicate process
regulated at multiple levels. A mishap at any point in the regulatory process can be
detrimental to organisms. On the same token, as long as all of the cellular processes that
contribute to the regulation of gene expression are in-check, then organisms are able to
maintain cellular homeostasis and overall health. The phrase “gene expression” is often
most closely associated with the regulation of translation. While this is not incorrect, we
must be aware that the regulation of gene expression begins on a much deeper level.
Even before the translation machinery begins translating an mRNA the mRNA itself is
subject to extensive regulation through the regulation of mRNA transcription and mRNA
degradation. Further, there are cellular quality control processes that ensure improperly
transcribed mRNAs are not further translated into deleterious or toxic proteins. These
quality control processes include: 1) no-go decay (NGD), which is responsible for
releasing ribosomes that have stalled on the mRNA and degrading the mRNA (Harigaya
and Parker, 2010), 2) nonstop decay (NSD), which degrades transcripts that lack a stop
codon allowing the ribosome to translate through the poly(A) tail adding a string of polylysines to the end of the protein (Frischmeyer et al., 2002), and 3) nonsense-mediated
mRNA decay (NMD), which is responsible for the rapid degradation of mRNAs that
harbor premature termination codons (PTCs; Akimitsu, 2008; Baker and Parker, 2004;
Behm-Ansmant et al., 2007; Chang et al., 2007; Frischmeyer and Dietz, 1999; Hentze
and Kulozik, 1999; Hilleren and Parker, 1999; Muhlemann et al., 2008; Schweingruber et
al., 2013; Shyu et al., 2008). While NMD was first discovered as an mRNA surveillance
220
mechanism it is now recognized that NMD plays a much broader role in the regulation of
wild-type mRNAs as well (Guan et al., 2006; He et al., 2003; Johansson et al., 2007;
Kalyna et al., 2012; Lelivelt and Culbertson, 1999; Mendell et al., 2004; Mitrovich and
Anderson, 2005; Peccarelli and Kebaara, 2014; Rayson et al., 2012; Rehwinkel et al.,
2005).
The known features that have been shown to target a wild-type mRNA for
degradation by NMD include: 1) a long 3’ UTR (Amrani et al., 2004; Kebaara and Atkin,
2009; Muhlrad and Parker, 1999), 2) translation of an upstream open reading frame
(uORF; Amrani et al., 2006; Barbosa et al., 2013; Gaba et al., 2005; Nyiko et al., 2009),
3) a start codon in a suboptimal context which can lead to leaky scanning and out of
frame initiation of translation (Welch and Jacobson, 1999), 4) the presence of
programmed ribosome frameshift (PRF) sites (Plant et al., 2004) and 5) the presence of
pre-mRNA introns and regulated alternative splicing resulting in PTCs (He et al., 1993;
Lewis et al., 2003; McGlincy and Smith, 2008; Ni et al., 2007).
Further, not all mRNAs with NMD-targeting signals are degraded by NMD. The
mechanisms of protection of mRNAs from degradation by NMD that have been
identified include: 1) inhibition of translation (Bertram et al., 2001; Czaplinski et al.,
1998; Gozalbo and Hohmann, 1990; He et al., 1993; Herrick et al., 1990; Keeling et al.,
2004; Kisselev et al., 2003; Peltz et al., 1992; Rospert et al., 2005; Tholstrup et al., 2012;
Zhang et al., 1997; Zuk and Jacobson, 1998), 2) stop codon readthrough (Bonetti et al.,
1995; Keeling et al., 2004; Namy et al., 2001), 3) translation reinitiation after a ribosome
encounters a stop codon (Hall and Thein, 1994; Neu-Yilik et al., 2011; Thein et al.,
221
1990), and, 4) trans-acting factor Pub1 (Ruiz-Echevarria and Peltz, 2000).
Although the NMD pathway has been studied extensively over the past decade,
and in multiple eukaryotes including Saccharomyces cerevisiae, Drosophila, C. elegans,
Arabidopsis, and mammalian cells, the exact molecular mechanism of the NMD pathway
remains obscure (Bedwell et al., 1997; Grimson et al., 2004; Hall and Thein, 1994; He et
al., 2003; Hentze and Kulozik, 1999; Kalyna et al., 2012; Kurihara et al., 2009; Maquat
and Carmichael, 2001; Maquat and Serin, 2001; Mendell et al., 2004; Pulak and
Anderson, 1993; Rayson et al., 2012; Rehwinkel et al., 2005). Because as many as one
third of all genetic diseases and cancers are linked to NMD, manipulation of the NMD
pathway has been a targeted area of study for the development of disease therapies
(Culbertson, 1999; Frischmeyer and Dietz, 1999; Kuzmiak and Maquat, 2006; Peltz et
al., 2013). However, the development of safe and effective therapies is hindered by the
obscurity of the exact molecular mechanisms of the NMD pathway. Thus, a thorough
understanding of how the cellular machinery distinguishes and wild-type mRNA from an
NMD substrate is a vital piece of being able to effectively treat NMD-related diseases.
Here we have provided a detailed investigation of a unique mRNA in
Saccharomyces cerevisiae, the SSY5 mRNA, which contains multiple NMD-targeting
signals but is not degraded by the NMD pathway. We show that many of the known
mechanisms that are responsible for the protection of mRNAs from NMD do not apply to
the SSY5 mRNA, thus, this mRNA is protected from NMD through a novel mechanism
(Chapter 2). We also show that the SSY5 mRNA is very tightly regulated during the
mRNA decay process (Chapter 3). Additionally, we also begin to characterize the
222
stability of two additional wild-type mRNAs in S. cerevisiae, the YAP1 and GCN4
mRNAs, which are protected from degradation by NMD despite containing translated
uORFs—an NMD-targeting signal (Chapter 4). Characterization of the stability of the
SSY5, YAP1 and GCN4 mRNAs provides valuable clues to understanding the molecular
mechanisms that control wild-type gene regulation by NMD.
Chapter 1: The S. cerevisiae SSY5 mRNA is not degraded by NMD despite multiple
NMD-targeting signals
The SSY5 mRNA was originally identified as a wild-type mRNA that should be
degraded by NMD in a screen for mRNAs in S. cerevisiae which contained long 3’ UTRs
(Kebaara and Atkin, 2009). Initial data from this study indicated that the SSY5 mRNA
was the only mRNA identified in the screen that contained a long 3’ UTR but showed no
difference in mRNA accumulation or half-life between wild-type and upf1Δ strains.
Upon closer analysis of the SSY5 mRNA we discovered that there were actually multiple
NMD-targeting signals present including: 1) a translated uORF, 2) a start codon in a poor
context that could lead to leaky scanning and out-of-frame initiation of translation, and 3)
the presence of five predicted ribosome frameshift sites. The long 3’ UTR of the SSY5
mRNA was previously characterized as a positive NMD-targeting signal (Obenoskey et
al., 2014). The translation of the uORF is shown with ribosome footprint data from
genome-wide studies that has been compiled in the GWIPS-viz genome browser (Michel
et al., 2014). However, further studies are needed to determine if leaky scanning or
programmed ribosome frameshifting is occurring.
223
Our studies confirm that the SSY5 mRNA shows similar accumulation and halflife in both wild-type and upf1Δ strains in two different S. cerevisiae genetic
backgrounds, BY4741 and W303. Additionally, differences in amino acid composition
of growth media does not affect SSY5 mRNA stability. The amino acid composition of
the growth media was taken into consideration because the SSY5 mRNA codes for an
essential proteolytic component of the SPS amino-acid-sensing complex in S. cerevisiae
(Conrad et al., 2014; Ljungdahl, 2009).
A previous study showed that when the long 3’ UTR of the SSY5 mRNA is placed
on an NMD-insensitive mRNA and the construct becomes a substrate for NMD
(Obenoskey et al., 2014). This data provides evidence that the long 3’ UTR of the SSY5
mRNA is sufficient to target an mRNA for degradation by NMD. Thus, there is a
mechanism that is protecting the SSY5 mRNA from being degraded by NMD. We further
showed that replacing the long 3’ UTR of the SSY5 mRNA with the short 3’ UTR of the
ADH1 mRNA does not affect the stability of the construct. So, the mechanism of
protection from NMD is not contained within the SSY5 mRNA 3’ UTR. We also
removed the SSY5 mRNA uORF by replacing the 5’ leader of the SSY5 mRNA with the
galactose promoter sequence. This construct did not have altered mRNA accumulation
or half-life in wild-type vs upf1Δ strains showing that the SSY5 mRNA uORF is not
required for protection from NMD.
The SSY5 mRNA is also not protected from NMD by known mechanism of
protection. We showed that the SSY5 mRNA co-fractionates with polyribosomes and is
actively translated. Additionally, although the SSY5 mRNA 3’ UTR is favorable for both
224
stop codon readthrough and reinitiation of translation, we do not find evidence of either.
Together, these studies show that the SSY5 mRNA is a very likely candidate for
degradation by the NMD pathway, and the protection of this mRNA from NMD occurs
through a novel mechanism. Future studies should investigate whether the NMD mRNP
is forming, either partially or completely, on the SSY5 mRNA. This would provide
information as to whether or not the SSY5 mRNA is recognized by the NMD machinery
as a substrate for NMD. Preliminary studies of this have begun but the experimental
procedures and controls need further optimization.
Additionally, we characterized the ENT4 mRNA in S. cerevisiae as a wild-type
mRNA that has a long 3’ UTR and is a substrate for NMD. This provides a positive
control for a wild-type substrate of NMD. Moreover, the current function of the gene
product of the ENT4 mRNA is unknown. Determining the function of the product of the
ENT4 mRNA will provide further insight into the physiological significance of wild-type
gene regulation by NMD in S. cerevisiae.
Further, the SPS-sensing pathway, which Ssy5 is an essential component of, is
critical for amino-acid sensing of the human fungal commensal and pathogen Candida
albicans. Importantly, the SPS pathway also controls the expression of virulence factors
in C. albicans (Davis et al., 2011). Further investigation of the stability of the SSY5
mRNA in C. albicans could provide important clues for understanding both the
pathogenicity of C. albicans in humans as well as SSY5 mRNA regulation by NMD.
225
Chapter 3: The SSY5 mRNA is very tightly regulated during mRNA degradation
Further characterization of the degradation of the SSY5 mRNA shows several
unique characteristics. First, the SSY5 mRNA does not follow a pattern representative of
wild-type mRNA decay through a deadenylation-dependent mechanism nor does the
mRNA follow a pattern of deadenylation-independent degradation representative of an
NMD substrate. Second, SSY5 mRNA accumulation is significantly increased when the
primary 5’3’ exonuclease Xrn1 is deleted. However, when decapping is blocked by
deletion of decapping component Dcp1, SSY5 mRNA accumulation is very modestly
increased. So, when the SSY5 mRNA is capped (in the dcp1Δ cells) the mRNA can still
be rapidly degraded 3’5’ by the exosome, but when the mRNA is decapped and 5’3’
degradation is blocked (in the xrn1Δ cells), 3’5’ degradation is not able to adequately
compensate. Further, deletion of any of the deadenylation components does not impact
SSY5 mRNA stability. Together these results show that SSY5 mRNA stability is very
tightly regulated at a step between mRNA decapping and 5’3’ exonucloelytic
degradation. This is fascinating because these processes occur subsequently and very
rapidly during mRNA decay. For completeness, future experiments should include an
analysis of the SSY5 mRNA in mutants of the components of the cytoplasmic exosome,
such as the catalytic component Dis3.
Additionally, analysis of the 3’ end of the SSY5 mRNA shows a pattern of 3’ end
shortening that is similar to what is seen for a wild-type mRNA that is deadenylated
before being decapped (PGK1 mRNA). Both the PGK1 and SSY5 mRNAs have longer
3’ ends in wild-type and upf1Δ strains and shorter 3’ ends in xrn1Δ and dcp1Δ strains.
226
This is in contrast to the ENT4 mRNA—a wild-type substrate degraded NMD in S.
cerevisiae—which shows the opposite pattern with a shorter 3’ end in wild-type and
upf1Δ strains and a longer 3’ end in xrn1Δ and dcp1Δ strains. This observation is likely
the result of the rate at which the 3’ end is degraded in wild-type versus NMD substrates
where 3’ end degradation occurs very rapidly following decapping for an NMD substrate
(Cao and Parker, 2003). However, to be able to conclude a change in length of the 3’ end
is the result of deadenylation we need to repeat the experiment with an RNase H control,
which removes the poly(A) tail.
An unexpected result of the studies in this section was that in a certain strain
background (not BY4741 or W303) the SSY5 mRNA does appear to be slightly stabilized
in the upf1Δ mutant. Initially, this was a troubling observation. However, upon further
consideration we have realized this can be used to our advantage. By determining the
difference(s) between this strain background and the BY4741 and W303 strains we can
drastically narrow down potential NMD-protecting mechanisms. Further characterization
of this strain background will be required. Once differences in the strain background are
identified we can begin studies to determine what might be, at least partially, responsible
for the protection of the SSY5 mRNA from NMD. This mechanism of protection may
then be further studied for other wild-type mRNAs that have NMD-targeting signals but
are not degraded by NMD, such as the YAP1 and GCN4 mRNAs.
227
Chapter 4: The S. cerevisiae wild-type mRNAs YAP1 and GCN4 also contain NMDtargeting signals but are not degraded by NMD
The YAP1 and GCN4 mRNAs were previously identified as wild-type mRNAs in
S. cerevisiae that have translated uORFS—an NMD-targeting signal—but are not
degraded by NMD (Ruiz-Echevarria et al., 1998; Vilela et al., 1998). A subsequent study
showed that both the YAP1 and GCN4 mRNAs are protected from degradation by NMD
due to the trans-acting factor Pub1 binding to a stabilizer element (STE) in the 5’ leader
of the mRNA downstream of the uORF stop codon (Ruiz-Echevarria and Peltz, 2000).
During our initial investigation into the protection of the SSY5 mRNA from NMD
we made the hypothesis that, based on the previous results, Pub1 could also be involved
in the protection of the SSY5 mRNA from NMD. Because Pub1 had already been clearly
implicated in the protection of both YAP1 and GCN4 mRNAs from NMD these were
included as controls. However, in our hands we were unable to reproduce the previously
published results despite extensive efforts. This means that we now have two additional
mRNAs in S. cerevisiae that have at least one NMD-targeting signal (we have not looked
for others) but are protected from degradation by NMD through an unknown mechanism.
We confirmed that both YAP1 and GCN4 mRNAs are protected from degradation
by NMD. However, half-life analysis in wild-type and upf1Δ strains provided stronger
confirmation of protection from NMD than steady-state mRNA accumulations. Because
the product of the GCN4 mRNA, Gcn4, is also involved in amino acid regulation we
again looked to see if the amino acid composition of the media had any influence on
mRNA stability of either of these mRNAs. We found that neither strain background nor
228
amino acid composition of the media influenced the stability of either mRNA. However,
as was the case when looking at SSY5 mRNA stability, the same holds true here where
we could not use a true amino-acid starvation media due to the auxotrophies of the strains
used. We could only provide fewer amino acids than are present in rich YAPD media.
True amino acid starvation or complete nitrogen starvation may influence mRNA
stability, but we were unable to test this.
We were unable to show that trans-acting factor Pub1 is solely responsible for the
protection of GCN4 and YAP1 mRNAs from NMD. However, we have preliminary data
that suggests, at least for the YAP1 mRNA, that Pub1 may be involved in the stability of
the mRNA but its role is more complex than previously indicated. Additionally, in
looking at the decay of the YAP1 and GCN4 mRNAs we find a unique pattern that does
not mimic that of a known NMD substrate (ENT4 mRNA), a wild-type mRNA degraded
by the deadenylation-dependent decapping pathway (PGK1 mRNA), or the SSY5 mRNA.
Unlike the SSY5 and ENT4 mRNAs, the YAP1 and GCN4 mRNAs show only a slight
increase in accumulation in an xrn1Δ strain, but still more accumulation than what is
observed for the PGK1 mRNA. Further, the YAP1 and GCN4 mRNAs show no
significant difference in accumulation in a dcp1Δ strain. This is again in contrast to the
SSY5 mRNA, which shows a slight increase in accumulation in a dcp1Δ strain. However,
it is similar to what is observed for the PGK1 mRNA, which shows no significant
difference in accumulation in a dcp1Δ strain. This does point to a tight window of
regulation between decapping and 5’3’ mRNA decay for both YAP1 and GCN4
mRNAs, but the fold changes are less than what are observed for the SSY5 mRNA, which
229
could suggest a larger role of 3’5’ mRNA decay for YAP1 and GCN4 mRNAs. The
deadenylation mutants also did not influence the accumulation of the YAP1 or GCN4
mRNAs consistent with what we observed for the SSY5, ENT4 and PGK1 mRNAs.
We also looked at the behavior of the 3’ end of the YAP1 mRNA in the wild-type,
upf1Δ, xrn1Δ and dcp1Δ strains. The data from this experiment shows that the 3’ end of
the YAP1 mRNA behaves similarly to the 3’ end of the PGK1 mRNA in all of the
mutants. The 3’ end of both YAP1 and PGK1 mRNAs is longer in wild-type and upf1Δ
strains and shorter in xrn1Δ and dcp1Δ strains. Again, to be able to conclude this change
in length is the result of deadenylation we need to repeat the experiment with an RNase H
control. This assay also needs to be performed for the GCN4 mRNA, which was not
originally included due to the cost of the kit and the minimal number of reactions
provided.
Final Thoughts
Although manipulation of the NMD pathway as a means of disease treatment
seems logical, and is currently being pursued all the way into clinical trials, this is
troubling given how much we still do not know about the underlying mechanisms of the
NMD pathway and wild-type gene regulation by NMD. Clinical trials of drugs, such as
Ataluren, are optimistically reporting no significant side effects of such therapies.
However, no long-term study data is available as these are newly developed therapies. It
is quite possible that side-effects may become apparent several years down the line.
Thus, it is absolutely critical that we continue attempts to elucidate the underlying
230
mechanisms of the NMD pathway in order to make treatments safer and more effective.
As with the development of any disease therapy, NMD-disease therapies will continue to
be modified as we learn more about the mechanisms of NMD. However, clinical trials of
NMD-related therapies at this point seems troublingly premature.
231
References
Akimitsu, N. (2008). Messenger RNA surveillance systems monitoring proper translation
termination. J Biochem 143, 1-8.
Amrani, N., Dong, S., He, F., Ganesan, R., Ghosh, S., Kervestin, S., Li, C., Mangus,
D.A., Spatrick, P., and Jacobson, A. (2006). Aberrant termination triggers nonsensemediated mRNA decay. Biochem Soc Trans 34, 39-42.
Amrani, N., Ganesan, R., Kervestin, S., Mangus, D.A., Ghosh, S., and Jacobson, A.
(2004). A faux 3'-UTR promotes aberrant termination and triggers nonsense-mediated
mRNA decay. Nature 432, 112-118.
Baker, K.E., and Parker, R. (2004). Nonsense-mediated mRNA decay: terminating
erroneous gene expression. Curr Opin Cell Biol 16, 293-299.
Barbosa, C., Peixeiro, I., and Romao, L. (2013). Gene expression regulation by upstream
open reading frames and human disease. PLoS Genet 9, e1003529.
Bedwell, D.M., Kaenjak, A., Benos, D.J., Bebok, Z., Bubien, J.K., Hong, J., Tousson, A.,
Clancy, J.P., and Sorscher, E.J. (1997). Suppression of a CFTR premature stop mutation
in a bronchial epithelial cell line. Nat Med 3, 1280-1284.
Behm-Ansmant, I., Gatfield, D., Rehwinkel, J., Hilgers, V., and Izaurralde, E. (2007). A
conserved role for cytoplasmic poly(A)-binding protein 1 (PABPC1) in nonsensemediated mRNA decay. EMBO J 26, 1591-1601.
Bertram, G., Innes, S., Minella, O., Richardson, J., and Stansfield, I. (2001). Endless
possibilities: translation termination and stop codon recognition. Microbiology 147, 255269.
Bonetti, B., Fu, L., Moon, J., and Bedwell, D.M. (1995). The efficiency of translation
termination is determined by a synergistic interplay between upstream and downstream
sequences in Saccharomyces cerevisiae. J Mol Biol 251, 334-345.
Cao, D., and Parker, R. (2003). Computational modeling and experimental analysis of
nonsense-mediated decay in yeast. Cell 113, 533-545.
Chang, Y.F., Imam, J.S., and Wilkinson, M.F. (2007). The nonsense-mediated decay
RNA surveillance pathway. Annu Rev Biochem 76, 51-74.
Conrad, M., Schothorst, J., Kankipati, H.N., Van Zeebroeck, G., Rubio-Texeira, M., and
Thevelein, J.M. (2014). Nutrient sensing and signaling in the yeast Saccharomyces
cerevisiae. FEMS Microbiol Rev 38, 254-299.
Culbertson, M.R. (1999). RNA surveillance. Unforeseen consequences for gene
expression, inherited genetic disorders and cancer. Trends Genet 15, 74-80.
232
Czaplinski, K., Ruiz-Echevarria, M.J., Paushkin, S.V., Han, X., Weng, Y., Perlick, H.A.,
Dietz, H.C., Ter-Avanesyan, M.D., and Peltz, S.W. (1998). The surveillance complex
interacts with the translation release factors to enhance termination and degrade aberrant
mRNAs. Genes Dev 12, 1665-1677.
Davis, M.M., Alvarez, F.J., Ryman, K., Holm, A.A., Ljungdahl, P.O., and Engstrom, Y.
(2011). Wild-type Drosophila melanogaster as a model host to analyze nitrogen source
dependent virulence of Candida albicans. PLoS One 6, e27434.
Frischmeyer, P.A., and Dietz, H.C. (1999). Nonsense-mediated mRNA decay in health
and disease. Hum Mol Genet 8, 1893-1900.
Frischmeyer, P.A., van Hoof, A., O'Donnell, K., Guerrerio, A.L., Parker, R., and Dietz,
H.C. (2002). An mRNA surveillance mechanism that eliminates transcripts lacking
termination codons. Science 295, 2258-2261.
Gaba, A., Jacobson, A., and Sachs, M.S. (2005). Ribosome occupancy of the yeast CPA1
upstream open reading frame termination codon modulates nonsense-mediated mRNA
decay. Mol Cell 20, 449-460.
Gozalbo, D., and Hohmann, S. (1990). Nonsense suppressors partially revert the decrease
of the mRNA level of a nonsense mutant allele in yeast. Curr Genet 17, 77-79.
Grimson, A., O'Connor, S., Newman, C.L., and Anderson, P. (2004). SMG-1 is a
phosphatidylinositol kinase-related protein kinase required for nonsense-mediated mRNA
Decay in Caenorhabditis elegans. Mol Cell Biol 24, 7483-7490.
Guan, Q., Zheng, W., Tang, S., Liu, X., Zinkel, R.A., Tsui, K.W., Yandell, B.S., and
Culbertson, M.R. (2006). Impact of nonsense-mediated mRNA decay on the global
expression profile of budding yeast. PLoS Genet 2, e203.
Hall, G.W., and Thein, S. (1994). Nonsense codon mutations in the terminal exon of the
beta-globin gene are not associated with a reduction in beta-mRNA accumulation: a
mechanism for the phenotype of dominant beta-thalassemia. Blood 83, 2031-2037.
Harigaya, Y., and Parker, R. (2010). No-go decay: a quality control mechanism for RNA
in translation. Wiley Interdiscip Rev RNA 1, 132-141.
He, F., Li, X., Spatrick, P., Casillo, R., Dong, S., and Jacobson, A. (2003). Genome-wide
analysis of mRNAs regulated by the nonsense-mediated and 5' to 3' mRNA decay
pathways in yeast. Mol Cell 12, 1439-1452.
He, F., Peltz, S.W., Donahue, J.L., Rosbash, M., and Jacobson, A. (1993). Stabilization
and ribosome association of unspliced pre-mRNAs in a yeast upf1- mutant. Proc Natl
Acad Sci U S A 90, 7034-7038.
Hentze, M.W., and Kulozik, A.E. (1999). A perfect message: RNA surveillance and
nonsense-mediated decay. Cell 96, 307-310.
233
Herrick, D., Parker, R., and Jacobson, A. (1990). Identification and comparison of stable
and unstable mRNAs in Saccharomyces cerevisiae. Mol Cell Biol 10, 2269-2284.
Hilleren, P., and Parker, R. (1999). Mechanisms of mRNA surveillance in eukaryotes.
Annu Rev Genet 33, 229-260.
Johansson, M.J., He, F., Spatrick, P., Li, C., and Jacobson, A. (2007). Association of
yeast Upf1p with direct substrates of the NMD pathway. Proc Natl Acad Sci U S A 104,
20872-20877.
Kalyna, M., Simpson, C.G., Syed, N.H., Lewandowska, D., Marquez, Y., Kusenda, B.,
Marshall, J., Fuller, J., Cardle, L., McNicol, J., et al. (2012). Alternative splicing and
nonsense-mediated decay modulate expression of important regulatory genes in
Arabidopsis. Nucleic Acids Res 40, 2454-2469.
Kebaara, B.W., and Atkin, A.L. (2009). Long 3'-UTRs target wild-type mRNAs for
nonsense-mediated mRNA decay in Saccharomyces cerevisiae. Nucleic Acids Res 37,
2771-2778.
Keeling, K.M., Lanier, J., Du, M., Salas-Marco, J., Gao, L., Kaenjak-Angeletti, A., and
Bedwell, D.M. (2004). Leaky termination at premature stop codons antagonizes
nonsense-mediated mRNA decay in S. cerevisiae. RNA 10, 691-703.
Kisselev, L., Ehrenberg, M., and Frolova, L. (2003). Termination of translation: interplay
of mRNA, rRNAs and release factors? EMBO J 22, 175-182.
Kurihara, Y., Matsui, A., Hanada, K., Kawashima, M., Ishida, J., Morosawa, T., Tanaka,
M., Kaminuma, E., Mochizuki, Y., Matsushima, A., et al. (2009). Genome-wide
suppression of aberrant mRNA-like noncoding RNAs by NMD in Arabidopsis. Proc Natl
Acad Sci U S A 106, 2453-2458.
Kuzmiak, H.A., and Maquat, L.E. (2006). Applying nonsense-mediated mRNA decay
research to the clinic: progress and challenges. Trends Mol Med 12, 306-316.
Lelivelt, M.J., and Culbertson, M.R. (1999). Yeast Upf proteins required for RNA
surveillance affect global expression of the yeast transcriptome. Mol Cell Biol 19, 67106719.
Lewis, B.P., Green, R.E., and Brenner, S.E. (2003). Evidence for the widespread
coupling of alternative splicing and nonsense-mediated mRNA decay in humans. Proc
Natl Acad Sci U S A 100, 189-192.
Ljungdahl, P.O. (2009). Amino-acid-induced signalling via the SPS-sensing pathway in
yeast. Biochem Soc Trans 37, 242-247.
Maquat, L.E., and Carmichael, G.G. (2001). Quality control of mRNA function. Cell 104,
173-176.
234
Maquat, L.E., and Serin, G. (2001). Nonsense-mediated mRNA decay: insights into
mechanism from the cellular abundance of human Upf1, Upf2, Upf3, and Upf3X
proteins. Cold Spring Harb Symp Quant Biol 66, 313-320.
McGlincy, N.J., and Smith, C.W. (2008). Alternative splicing resulting in nonsensemediated mRNA decay: what is the meaning of nonsense? Trends Biochem Sci 33, 385393.
Mendell, J.T., Sharifi, N.A., Meyers, J.L., Martinez-Murillo, F., and Dietz, H.C. (2004).
Nonsense surveillance regulates expression of diverse classes of mammalian transcripts
and mutes genomic noise. Nat Genet 36, 1073-1078.
Michel, A.M., Fox, G., A, M.K., De Bo, C., O'Connor, P.B., Heaphy, S.M., Mullan, J.P.,
Donohue, C.A., Higgins, D.G., and Baranov, P.V. (2014). GWIPS-viz: development of a
ribo-seq genome browser. Nucleic Acids Res 42, D859-864.
Mitrovich, Q.M., and Anderson, P. (2005). mRNA surveillance of expressed
pseudogenes in C. elegans. Curr Biol 15, 963-967.
Muhlemann, O., Eberle, A.B., Stalder, L., and Zamudio Orozco, R. (2008). Recognition
and elimination of nonsense mRNA. Biochim Biophys Acta 1779, 538-549.
Muhlrad, D., and Parker, R. (1999). Aberrant mRNAs with extended 3' UTRs are
substrates for rapid degradation by mRNA surveillance. RNA 5, 1299-1307.
Namy, O., Hatin, I., and Rousset, J.P. (2001). Impact of the six nucleotides downstream
of the stop codon on translation termination. EMBO Rep 2, 787-793.
Neu-Yilik, G., Amthor, B., Gehring, N.H., Bahri, S., Paidassi, H., Hentze, M.W., and
Kulozik, A.E. (2011). Mechanism of escape from nonsense-mediated mRNA decay of
human beta-globin transcripts with nonsense mutations in the first exon. RNA 17, 843854.
Ni, J.Z., Grate, L., Donohue, J.P., Preston, C., Nobida, N., O'Brien, G., Shiue, L., Clark,
T.A., Blume, J.E., and Ares, M., Jr. (2007). Ultraconserved elements are associated with
homeostatic control of splicing regulators by alternative splicing and nonsense-mediated
decay. Genes Dev 21, 708-718.
Nyiko, T., Sonkoly, B., Merai, Z., Benkovics, A.H., and Silhavy, D. (2009). Plant
upstream ORFs can trigger nonsense-mediated mRNA decay in a size-dependent manner.
Plant Mol Biol 71, 367-378.
Obenoskey, J., Lane, D.R., Atkin, A.L., and Kebaara, B.W. (2014). Immunity of the
Saccharomyces cerevisiae SSY5 mRNA to nonsense-mediated mRNA decay. Front Mol
Biosci 1, 25.
Peccarelli, M., and Kebaara, B.W. (2014). Regulation of natural mRNAs by the
nonsense-mediated mRNA decay pathway. Eukaryot Cell 13, 1126-1135.
235
Peltz, S.W., Donahue, J.L., and Jacobson, A. (1992). A mutation in the tRNA
nucleotidyltransferase gene promotes stabilization of mRNAs in Saccharomyces
cerevisiae. Mol Cell Biol 12, 5778-5784.
Peltz, S.W., Morsy, M., Welch, E.M., and Jacobson, A. (2013). Ataluren as an agent for
therapeutic nonsense suppression. Annu Rev Med 64, 407-425.
Plant, E.P., Wang, P., Jacobs, J.L., and Dinman, J.D. (2004). A programmed -1 ribosomal
frameshift signal can function as a cis-acting mRNA destabilizing element. Nucleic Acids
Res 32, 784-790.
Pulak, R., and Anderson, P. (1993). mRNA surveillance by the Caenorhabditis elegans
smg genes. Genes Dev 7, 1885-1897.
Rayson, S., Arciga-Reyes, L., Wootton, L., De Torres Zabala, M., Truman, W., Graham,
N., Grant, M., and Davies, B. (2012). A role for nonsense-mediated mRNA decay in
plants: pathogen responses are induced in Arabidopsis thaliana NMD mutants. PLoS One
7, e31917.
Rehwinkel, J., Letunic, I., Raes, J., Bork, P., and Izaurralde, E. (2005). Nonsensemediated mRNA decay factors act in concert to regulate common mRNA targets. RNA
11, 1530-1544.
Rospert, S., Rakwalska, M., and Dubaquie, Y. (2005). Polypeptide chain termination and
stop codon readthrough on eukaryotic ribosomes. Rev Physiol Biochem Pharmacol 155,
1-30.
Ruiz-Echevarria, M.J., Gonzalez, C.I., and Peltz, S.W. (1998). Identifying the right stop:
determining how the surveillance complex recognizes and degrades an aberrant mRNA.
EMBO J 17, 575-589.
Ruiz-Echevarria, M.J., and Peltz, S.W. (2000). The RNA binding protein Pub1 modulates
the stability of transcripts containing upstream open reading frames. Cell 101, 741-751.
Schweingruber, C., Rufener, S.C., Zund, D., Yamashita, A., and Muhlemann, O. (2013).
Nonsense-mediated mRNA decay - mechanisms of substrate mRNA recognition and
degradation in mammalian cells. Biochim Biophys Acta 1829, 612-623.
Shyu, A.B., Wilkinson, M.F., and van Hoof, A. (2008). Messenger RNA regulation: to
translate or to degrade. EMBO J 27, 471-481.
Thein, S.L., Hesketh, C., Taylor, P., Temperley, I.J., Hutchinson, R.M., Old, J.M., Wood,
W.G., Clegg, J.B., and Weatherall, D.J. (1990). Molecular basis for dominantly inherited
inclusion body beta-thalassemia. Proc Natl Acad Sci U S A 87, 3924-3928.
Tholstrup, J., Oddershede, L.B., and Sorensen, M.A. (2012). mRNA pseudoknot
structures can act as ribosomal roadblocks. Nucleic Acids Res 40, 303-313.
236
Vilela, C., Linz, B., Rodrigues-Pousada, C., and McCarthy, J.E. (1998). The yeast
transcription factor genes YAP1 and YAP2 are subject to differential control at the levels
of both translation and mRNA stability. Nucleic Acids Res 26, 1150-1159.
Welch, E.M., and Jacobson, A. (1999). An internal open reading frame triggers nonsensemediated decay of the yeast SPT10 mRNA. EMBO J 18, 6134-6145.
Zhang, S., Welch, E.M., Hogan, K., Brown, A.H., Peltz, S.W., and Jacobson, A. (1997).
Polysome-associated mRNAs are substrates for the nonsense-mediated mRNA decay
pathway in Saccharomyces cerevisiae. RNA 3, 234-244.
Zuk, D., and Jacobson, A. (1998). A single amino acid substitution in yeast eIF-5A
results in mRNA stabilization. EMBO J 17, 2914-2925.